Fabrication of vascularized tissue using microfabricated two-dimensional molds

ABSTRACT

Methods and materials for making complex, living, vascularized tissues for organ and tissue replacement, especially complex and/or thick, structures, such as liver tissue is provided. Tissue lamina is made in a system comprising an apparatus having (a) a first mold or polymer scaffold, a semi-permeable membrane, and a second mold or polymer scaffold, wherein the semi-permeable membrane is disposed between the first and second molds or polymer scaffolds, wherein the first and second molds or polymer scaffolds have means defining microchannels positioned toward the semi-permeable membrane, wherein the first and second molds or polymer scaffolds are fastened together; and (b) animal cells. Methods for producing complex, three-dimensional tissues or organs from tissue lamina are also provided.

RELATED APPLICATIONS

This application is a continuation of U.S. Ser. No. 14/153,591, filedJan. 13, 2014, which is a continuation of U.S. Ser. No. 12/785,865,filed May 24, 2010, now U.S. Pat. No. 8,642,336 which was a divisionalof U.S. Ser. No. 10/187,247, filed Jun. 28, 2002, now U.S. Pat. No.7,759,113. This application also claims the benefit of U.S. Ser. No.60/367,675, filed Mar. 25, 2002.

STATEMENT OF POTENTIAL GOVERNMENT INTEREST

This invention was made with Government support under Grant NumberHL6042404 awarded by the National Institutes of Health as well asDAMD17-99-2-9001 and DAMD17-02-0006 from the Department of the Army. TheGovernment has certain rights in this invention.

Each of the foregoing applications and patents and articles, and eachdocument cited or referenced in each of the foregoing applications andpatents and articles, including during the prosecution of each of theforegoing applications and patents (“application and article citeddocuments”), and any manufacturer's instructions or catalogues for anyproducts cited or mentioned in each of the foregoing applications andpatents and articles and in any of the application and article citeddocuments, are hereby incorporated herein by reference. Furthermore, alldocuments cited in this text, and all documents cited or referenced indocuments cited in this text, and any manufacturer's instructions orcatalogues for any products cited or mentioned in this text or in anydocument hereby incorporated into this text, are hereby incorporatedherein by reference. Documents incorporated by reference into this textor any teachings therein can be used in the practice of this invention.Documents incorporated by reference into this text are not admitted tobe prior art. Furthermore, authors or inventors on documentsincorporated by reference into this text are not to be considered to be“another” or “others” as to the present inventive entity and vice versa,especially where one or more authors or inventors on documentsincorporated by reference into this text are an inventor or inventorsnamed in the present inventive entity.

FIELD OF THE INVENTION

The present invention generally relates to the fields of organtransplantation and reconstructive surgery, and to the new field oftissue engineering. It more specifically is a new method and materialsfor generating tissues requiring a blood vessel supply and other complexcomponents such as a nerve supply, drainage system and/or lymphaticsystem.

BACKGROUND

Vital organ failure is one of the most critical problems facing thehealth care field today. Organ transplantation, as currently practiced,has become a major lifesaving therapy for patients afflicted withdiseases that destroy vital organs including the heart, liver, lungs,kidney and intestine. However, the shortage of organs needed fortransplantation has become critical and continues to worsen. Forexample, in the United States, the number of patients awaiting an organfor transplant has risen above 75,000. Despite advances in living donororgan transplantation, a severe shortage of donor organs available tothese patients remains as the crux of the problem. Likewise, every majorfield of reconstructive surgery reaches the same barrier of tissueshortage. Orthopedic surgery, vascular surgery, cardiac surgery, generalsurgery, neurosurgery, and the others all share this fundamentalproblem. Therefore, countless patients suffer as a result. Mechanicaldevices provide one approach to addressing the organ and tissueshortage. Xenografts provide another approach. However, due to theintrinsic limitations of these technologies, these approaches are onlypartial solutions to the problem.

Over the last several years, the new field of tissue engineering hasarisen to meet this need. The field brings the expertise of physicians,life scientists and engineers together to solve problems of generatingnew tissues for transplantation and surgical reconstruction. Tissueengineering can be a complete and permanent solution to the problem oforgan loss or failure, but the primary challenge for tissue engineeringvital organs is the requirement for a vascular supply for nutrient andmetabolite transfer. The initial approaches to this problem weredescribed in the 1980's. Yannas, et al., Science 221, 1052 (1981) andBurke, et al., Ann Surg 194, 413 (1981) relate to methods to generatenew tissues in vivo by implanting non-living materials such as modifiedcollagens which are seeded with cells to promote guided regeneration oftissue such as skin. Langer, et al. Science 260, 920 (1993) and Vacanti,et al., Materials Research Society 252, 367 (1992) involve syntheticfibrous matrices to which tissue specific cells were added in vitro. Thematrices are highly porous and allow mass transfer to the cells in vitroand after implantation in vivo. After implantation, new blood vesselsgrow into the devices to generate a new vascularized tissue. However,the relatively long time course for angiogenesis limits the size of thenewly formed tissue.

The field of Tissue Engineering is undergoing explosive growth. See, forexample, Vacanti, et al., Lancet 354, 32 (1999); Langer, et al, Science260, 920 (1993); Rennie, J. Scientific American 280, 37 (1999); andLysaght, et al., Tissue Eng 4, 231 (1998). Virtually every tissue andorgan of the body has been studied; many tissue-engineering technologiesare becoming available. See Lysaght, et al. Tissue Eng 4, 231 (1998);Bell, et al., Science 221, 1052 (1981); Burke, et al., Ann Surg 194, 413(1981); Compton, et al., Laboratory Investigation 60, 600 (1989);Parenteau, et al., Journal of Cellular Biochemistry 45, 24 (1991);Parenteau, et al., Biotechnology and Bioengineering 52, 3 (1996);Purdue, et al., J. Burn Care Rehab 18, 52 (1997); Hansbrough and Franco,Clinical Plastic Surg 25, 407 (1998); Vacanti, et al., MaterialsResearch Society 252, 367 (1992).

Over time, several techniques to engineer new living tissue have beenstudied. Technologies include the use of growth factors to stimulatewound repair and regeneration, techniques of guided tissue regenerationusing non-living matrices to guide new tissue development, celltransplantation, and cell transplantation on matrices. More recently,new understanding in stem cell biology has led to studies of populationsof primordial cells, stem cells, or embryonic stem cells to use intissue engineering approaches.

In parallel to these advances, the rapidly emerging field ofMicroElectroMechanical Systems (MEMS) has penetrated a wide array ofapplications, in areas as diverse as automotives, inertial guidance andnavigation, microoptics, chemical and biological sensing, and, mostrecently, biomedical engineering, McWhorter, et al. “Micromachining andTrends for the Twenty-First Century”, in Handbook of Microlithography,Micromachining and Microfabrication, ed. P. Rai-Choudhury, (Bellingham,Wash.: SPIE Press, 1997). Microfabrication technology has been used inimportant studies in cell and developmental biology to understandcomplex biologic signaling events occurring at the cell membrane-surfaceinterface, as described, for example, by Kane, et al., Biomaterials 20,2363 (1999). It has also been used in tissue engineering to guide cellbehavior and the formation of small units of tissue, as described byGriffith, et al., Annals of Blamed. Eng., 26 (1998).

Microfabrication methods for MEMS represent an extension ofsemiconductor wafer process technology originally developed for theintegrated circuit (IC) industry. Control of features down to thesubmicron level is routinely achieved in IC processing of electricalcircuit elements; MEMS technology translates this level of control intomechanical structures at length scales stretching from less than 1micron (μm) to greater than 1 centimeter (cm). Standard bulkmicromachining enables patterns of arbitrary geometry to be imprintedinto wafers using a series of subtractive etching methods.Three-dimensional structures can be realized by superposition of theseprocess steps using precise alignment techniques. Several groups(Griffith, et al., Annals of Biomed. Eng., 26 (1998); Folch, et al.,Biotechnology Progress, 14, 388 (1998)) have used these highly precisesilicon arrays to control cell behavior and study gene expression andcell surface interactions. However, this approach is essentially atwo-dimensional technology and it is unknown whether it can be adaptedto the generation of thick, three-dimensional tissues.

PCT US96/09344 by Massachusetts Institute of Technology involves athree-dimensional printing process, a form of solid free formfabrication, which builds three-dimensional objects as a series oflayers. This process uses polymer powders in layers bound by polymerbinders whose geometry is dictated by computer-assisted design andmanufacture. This technique allows defined internal architectures, whichcould include branching arrays of channels mimicking a vascular supply.However, this technique is limited by the characteristics and chemistryof the particular polymers. Also, it severely limits the types of tissueto be fabricated. For example, these polymer walls do not allow theplasma exchange that is needed in the alveolar capillary wall of thelung.

A further limitation of the prior art methods of tissue engineering isrelated to mass transport. Cells must be within approximately 100 μm ofa capillary blood supply. Tissue engineered constructs without a bloodsupply develop hypoxia and nutrient deprivation. Without vasculature,cells in constructs larger than 1-2 mm experience significant necrosis.To date, all approaches in tissue engineering have relied on thein-growth of blood vessels into tissue-engineered devices to achievepermanent vascularization. This strategy has worked well for manytissues; however, it falls short for thick, complex tissues such aslarge vital organs, including liver, kidney, and heart. See Eiselt, etal., Biotechnol. Prog. 14, 134 (1998). Novel methods and devices thatenable the production of thick, complex tissue-engineered structureswould be highly desirable.

OBJECTS AND SUMMARY OF THE INVENTION

To overcome obstacles known in the art, an approach to provide thickstructures with preexisting vasculature before implantation wasdeveloped using microfabrication techniques, such as three-dimensionalprinting to provide an ordered array of branching channels in asubstrate formed of a suitable material, such as silicon or abiocompatible polymer, which are then seeded with cells. A completebranching vascular circulation is made in two dimensions on the surfaceof the material using microfabrication. The two-dimensional structurecan then be lifted or otherwise separated from the silicon mold andfolded or rolled into a compact three-dimensional structure.

The laminated tissue structures comprise multiple layers wherein eachlayer comprises tissue and vasculature assembled adjacent to each otherby folding and compacting. The vasculature is in three dimensionsthroughout the structure and the structure optionally has connectionsfor flow into and out of the vasculature. The laminated tissuestructures can be implanted directly by connecting blood vessels to flowinto and out of the vasculature. As such, the present inventionovercomes problems known in the art of tissue engineering, which islimited to the production of very thin structures.

An object of the present invention can be to provide a method andmaterials for making complex, living, vascularized tissues for organ andtissue replacement, especially complex and/or thick structures, such asliver tissue.

The invention provides an apparatus for making tissue lamina comprisinga first mold or polymer scaffold, a semi-permeable membrane, and asecond mold or polymer scaffold, wherein the semi-permeable membrane isdisposed between the first and second molds and/or polymer scaffolds,wherein the first and second molds and/or polymer scaffolds havemicrochannels or compartments positioned toward the semi-permeablemembrane, wherein the first and second molds and/or polymer scaffoldsare fastened together, and wherein each of the first and second moldsand/or polymer scaffolds and the semi-permeable membrane are comprisedof material that is suitable for attachment and culturing of animalcells. The semi-permeable membrane enables transport of small molecules,such as oxygen, through thin bulk layers, but does not require actualpores for transport.

The apparatuses of the invention are made from layers comprising atleast a first mold or polymer scaffold, a second mold or polymerscaffold and a semi-permeable membrane. These layers can be assembledsuch that they are in complete registration (i.e. perfectly aligned), orin partial registration (i.e. imperfectly aligned).

The apparatus can optionally be in fluid communication with nutrientsupply and excretion removal lines for culturing animal cells, and canfurther comprise a pumping means for circulating fluid throughout theapparatus. The direction of flow can be controlled or directed asneeded, and can be in all directions. The pumping means can comprise asyringe, a peristaltic pump or any other pumping means known in the artof cell or tissue culture.

The semi-permeable membrane allows gas exchange, diffusion of nutrients,and waste removal. The first mold or polymer scaffold can comprise thecirculation through which blood, plasma or media with appropriate levelsof oxygen can be continuously circulated to nourish the cells/tissue inone or more additional molds and/or polymer scaffolds. The second moldor polymer scaffold can comprise a reservoir for the functional cells ofan organ, and optionally includes inlets for neural inervation or otheractivity.

The invention further comprises a system for making tissue laminacomprising (a) an apparatus for making tissue lamina comprising a firstmold or polymer scaffold, a semi-permeable membrane, and a second moldor polymer scaffold, wherein the semi-permeable membrane is disposedbetween the first and second molds and/or polymer scaffolds, wherein thefirst and second molds and/or polymer scaffolds have means definingmicrochannels positioned toward the semi-permeable membrane, wherein thefirst and second molds and/or polymer scaffolds are fastened together,and wherein each of the first and second molds and/or polymer scaffoldsand the semi-permeable membrane are comprised of material that issuitable for attachment and culturing of animal cells; and (b) animalcells.

The invention also provides a method of making an apparatus for makingtissue lamina comprising the steps of (a) positioning a semi-permeablemembrane between a first and second mold and/or polymer scaffold,wherein the first and second molds and/or polymer scaffolds have meansdefining microchannels positioned toward the semi-permeable membrane andwherein each of the first and second molds and/or polymer scaffolds andthe semi-permeable membrane are comprised of material that is suitablefor attachment and culturing of animal cells; and (b) fastening thefirst and second molds and/or polymer scaffolds together such that thesemi-permeable membrane is disposed between the molds and/or polymerscaffolds.

As described herein, complex tissues are formed by laminating layers ofthin vascularized tissues to form thick tissue structures or morecomplex organ equivalents. The thin vascularized layers of tissue laminaare formed by:

(a) positioning a semi-permeable membrane between a first and secondmold and/or polymer scaffold, wherein the first and second molds and/orpolymer scaffolds have means defining microchannels positioned towardthe semi-permeable membrane, and wherein each of the first and secondmolds and/or polymer scaffolds and the semi-permeable membrane arecomprised of material that is suitable for attachment and culturing ofanimal cells;

(b) fastening the first and second molds and/or polymer scaffoldstogether such that the semi-permeable membrane is disposed between themolds and/or polymer scaffolds, thereby forming an apparatus; and

(c) culturing cells in the microchannels of the molds and/or polymerscaffolds. Culturing cells in microchannels can comprise

-   -   (i) seeding animal cells into the microchannels of the first        mold or polymer scaffold;    -   (ii) culturing the animal cells of (i) under conditions such        that they form blood vessels or other lumenal structures;    -   (iii) seeding animal cells into the microchannels of the second        mold or polymer scaffold; and    -   (iv) culturing the animal cells of (ii) under conditions such        that they form parenchymal tissue.

The apparatus and tissues therein can optionally be connected in fluidcommunication with nutrient supply and excretion removal lines forculturing animal cells, and fluid can further be circulated through theapparatus by means of a pump. The pump can comprise a syringe, aperistaltic pump or any other pumping means known in the art of cell ortissue culture.

Three dimensional tissue can be formed by gently lifting tissue from themold and/or polymer scaffold using techniques such as fluid flow andother supporting material, as necessary. Where the polymer scaffold iscomprised of biodegradable monomers, hydrolysis of the monomers resultsin degradation of the scaffold over time.

The tissue can be systematically folded or rolled and compacted into athree-dimensional vascularized structure. This structure can beimplanted into animals or patients by directly connecting the bloodvessels. Optionally, the semi-permeable membrane can be removed.Immediate perfusion of oxygenated blood occurs, which allows survivaland function of the entire living mass.

The two-dimensional surface of the mold can also be varied to aid in thefolding and compacting process. For example, the surface can be changedfrom planar to folded in an accordion-like fashion. It can be stackedinto multiple converging plates. It can be curvilinear or have multipleprojections.

Alternatively, multiple molds and/or polymer scaffolds can be stackedadjacent to one another, making multiple vascularized layers of tissuelamina until the desired complex structure (e.g. organ equivalent) isformed. These structures can then be implanted and optionally, thevasculature anastomized into the existing vasculature to provide animmediate blood supply for the implanted organ equivalent.Alternatively, these structures comprise extracorporeal support devices.

A preferred method of making multiple layers of tissue lamina in threedimensions is to form an apparatus by positioning a semi-permeablemembrane between two molds and/or polymer scaffolds, which molds havemicrochannels positioned toward the semi-permeable membrane, and whereinthe molds and/or polymer scaffolds and the semi-permeable membranecomprise material that is suitable for attachment and culturing ofanimal cells. The molds and/or polymer scaffolds are fastened togethersuch that the semi-permeable membrane is between them, Cells are thencultured in the microchannels of the molds and/or polymer scaffolds toform tissue lamina, which is removed from the apparatus and folded orrolled to form multiple layers of tissue lamina in three dimensions.

In another embodiment, multiple layers of tissue lamina in threedimensions are created by forming an apparatus from two molds and/orpolymer scaffolds and a semi-permeable membrane, as described above, andfastening a third mold or polymer scaffold adjacent to the first orsecond mold or polymer scaffold. The third mold or polymer scaffold alsohas microchannels, which are positioned toward the adjacent mold orpolymer scaffold. A second semi-permeable membrane can separate thethird mold or polymer scaffold from the adjacent mold or polymerscaffold. Microchannels in successive layers can be connected by throughholes to create a parallel channel network in three dimensions. Cellsare cultured in the microchannels of the first, second and third moldsand/or polymer scaffolds to form three dimensional tissue. Additionalmold and/or polymer scaffold layers and optionally, semi-permeablemembranes, can be added to create thicker tissue.

The systems and methods of the invention can be implanted into a subjectto supplement or replace the biological function of a tissue or organ.Alternatively, the systems and methods can remain ex vivo, serving asextracorporeal devices to supplement or replace biological function.

Examples of tissues and organs which can be fabricated using thesemethods include, but are not restricted to, organs currentlytransplanted such as heart, liver, pancreas, lung, kidney and intestine.Other tissues such as muscle, bone, breast, reproductive and neuraltissue could also be engineered.

These and other embodiments are disclosed or are obvious from andencompassed by, the following Detailed Description.

BRIEF DESCRIPTION OF THE DRAWINGS

The following Detailed Description, given by way of example, but notintended to limit the invention to specific embodiments described, maybe understood in conjunction with the accompanying Figures, incorporatedherein by reference, in which:

FIG. 1A shows a silicon wafer with a network of microchannels.

FIG. 1B shows a polymer scaffold with a network of microchannels.

FIG. 2 shows a schematic diagram of one method for making a polymerscaffold from a micromachined silicon wafer using MEMS replica molding.

FIG. 3 shows a schematic of a process for fabricating U-shaped trenchesto make a branching pattern on silicon wafers.

FIG. 4 shows a more detailed schematic describing a process for theproduction of a complex structure comprising channels of varying depths.

FIG. 5 shows a schematic of a pattern etched using aninductively-coupled plasma (ICP) system.

FIG. 6 shows a schematic of an etched surface showing a branchingstructure that branches out from a single inlet and then converges backinto a single outlet.

FIGS. 7A, B, and C show schematics of a cross-sectional view ofdifferent etched channels in the surface of FIG. 6.

FIGS. 8A, B and C show schematics of a process for making a tissuelayer.

FIGS. 9A and 9B show schematic diagrams of a cross section of anapparatus for tissue engineering and artificial organ support. Theapparatus in FIG. 9A comprises a compartment for circulatory flow (1), asemi-permeable membrane for mass transfer of oxygen, nutrients and waste(2), and a compartment for functional cells and excretory system. FIG.9B shows the apparatus of 9A seeded with vascular cells or cells thatform lumen (e.g. biliary ducts) (4) and functional cells (e.g.hepatocytes) (5).

FIGS. 10A-10C show electron micrographs of a semi-permeable membranemade using the TIPS procedure. FIGS. 10A and 10B show the membranesurface at 650× and 6500× magnification, respectively. FIG. 10C showsthe membrane in cross-section at 650× magnification.

FIG. 11 shows a schematic top drawing of a mold or polymer scaffold. Thetriangles represent areas coated with cell adhesion molecules to promotethe adhesion of cells (e.g. hepatocytes). The white areas between thetriangles represent microchannels; in some applications, they are notcoated with cell adhesion molecules, and so are open for colonization bycells that can form vascular tissue (e.g. endothelial cells). The blackcircle in the middle of each hexagon is a vertical through-hole.

FIGS. 12A-G show schematics of various surfaces and how tissue layersmight be assembled from them.

FIG. 13 shows a schematic of an assembled complex tissue or organ formedby the process of FIG. 8.

FIG. 14 shows how the organ of FIG. 5 can be connected to a fluid byanastomosis of the inlet and outlet.

FIG. 15 shows a set of bar graphs demonstrating continued albuminproduction by hepatocyte cells cultured in a polymer scaffold of theinvention. Albumin concentration in culture medium was measured every 24hours for 5 days pre-cell detachment using an enzyme linkedimmunosorbent assay. No significant differences were observed betweenday 2, day 3, and day 4 (p<0.05 by the paired t-test).

FIG. 16A shows a sample vascular branching network pattern used forsilicon and pyrex wafer micromachining.

FIG. 16B shows the optical micrograph or portion of the capillarynetwork etched into the silicon wafer using the process shown in FIG. 3.

FIG. 16C shows a scanning electron micrograph of an anisotrophic etchingprocess used to form angled sidewall trenches.

FIGS. 17A-C show phase-contrast photographs of small hepatocytes andnonparenchymal cells cultured on regular culture flasks. FIG. 17A showscells in culture at Day 3. FIG. 17B shows cells in culture at Day 5.FIG. 17C shows cells in culture at Day 10. Scale bar, 100 μm (originalmagnification ×100).

FIGS. 18A and 18B show a cell sheet lifted from a silicon wafer. FIG.18A shows macroscopic appearance and FIG. 18B shows microscopicappearance (original magnification ×30).

FIG. 19 shows albumin production by small hepatocytes at day 3, 5, 7,and 10 (μg/day).

FIGS. 20A-D show H & E staining of implanted constructs. FIG. 20A showsconstructs at 2 weeks. Arrows indicate bile ductular structures. FIG.20B shows constructs at 1 month. Arrows indicate bile ductularstructures. FIG. 20C shows constructs at 2 months. The large clusters ofhepatocytes over five cell layers thick were observed at 1 and 2 months.FIG. 20D shows constructs at 1 month. The implanted construct wasoccupied by the bile ductular structures.

FIGS. 21A-D show immunohistochemical staining of implanted constructs.FIG. 21A shows pan-cytokeratin staining at 1 month. FIG. 21B showsalbumin staining at 1 month. Arrows indicate bile ductular structures.FIG. 21C shows transferrin staining at 1 month. Arrows indicate bileductular structures. FIG. 21D shows GGT staining at 1 month. Arrowsindicate bile ductular structures with luminal staining. Arrow headsindicate slightly stained hepatocytes.

FIG. 22 shows H & E staining at 1 month. Arrow indicates the bileductular structure composed of both biliary epithelial cells and ahepatocyte. Arrow heads indicate the bile ductular structures composedof biliary epithelial cells.

FIGS. 23A-B show transmission electron microscopy (TEM) of an implantedconstruct. FIG. 23A is at magnification (×2500). FIG. 23B is at highmagnification (×15000).

FIG. 24 shows the area occupied by implanted constructs (μg²/section).Total area and bile ducts area are expressed as mean+/−SD.

FIG. 25 shows a schematic diagram of a micromachined apparatus fortissue engineered renal replacement. The apparatus comprises acompartment with a glomerular endothelial filter for circulatory flow(42), a semi-permeable membrane for mass transfer of oxygen, nutrientsand waste (44), and a compartment with a proximal tubule networkexcretory system, which includes inlets for filtration of urine (46).

FIG. 26 shows a cross section of a micromachined apparatus for tissueengineered renal replacement. The apparatus comprises a compartment witha glomerular endothelial filter for circulatory flow, a semi-permeablemembrane for mass transfer of oxygen, nutrients and waste, and acompartment with a proximal tubule network excretory system, whichincludes inlets for filtration of urine. Each compartmentalized layer ofthe apparatus comprises a biocompatible polymer and the layers areseparated by a semi-permeable membrane comprising a microporous polymer.

FIG. 27 shows a cross section of a micromachined apparatus for tissueengineered renal replacement. The direction of flow of glomerularultrafiltrate is shown. Flow originates in the layer comprisingglomerular endothelium, passes through the semi-permeable membrane tolayer comprising the proximal tubule network where reabsorption occurs.

FIG. 28 shows a cross section of a micromachined apparatus for tissueengineered renal replacement comprising multiple stacked layers. Theapparatus comprises repeating, stacked units, each unit comprising acompartment with a glomerular endothelial filter for circulatory flow, asemi-permeable membrane for mass transfer of oxygen, nutrients andwaste, and a compartment with a proximal tubule network excretorysystem, which includes inlets for filtration of urine.

FIG. 29 shows human microvascular cells at 14 days after seeding inmicrochannels.

FIG. 30 shows proximal tubule cells growing in a poly dimethyl-siloxane(PDMS) polymer scaffold at approximately 5 hours after seeding.

FIG. 31 shows proximal tubule cells growing in a poly dimethyl-siloxane(PDMS) polymer scaffold at 2 days after seeding.

FIG. 32 shows proximal tubule cells growing in a poly dimethyl-siloxane(PDMS) polymer scaffold at 6 days after seeding.

DETAILED DESCRIPTION

The invention provides for the construction of structures comprisingtissue in thick layers; enabling fabrication of an entire organ, orportion thereof, having sufficient oxygen transport, nutrient andmetabolite movement. The methods of the invention employ a new approachfor fabricating three-dimensional vascularized tissues fortransplantation in human recipients in need of vital organs and othertissues requiring a blood supply. A two-dimensional (x, y) mold isfabricated using high-resolution molding processes, such asmicromachined wafer technology, thick photoresist processes, or othertechniques, to create a patterned of micromachined, small dimensionedchannels (“microchannels”), such that the micromachined channels areconnected for the circulation of fluid in the multilayer apparatus.Microchannels can comprise, for example, open-faced channels defined bywalls extending from a tissue-defining surface into a substrate. Theinvention also encompasses a substrate wherein the tissue-definingsurface comprises an open-faced compartment defined by walls extendingfrom a tissue-defining surface into a substrate.

Thus, the invention provides low-cost, scalable techniques for producingorgans, or portions thereof, large enough to transplant into a subject,such as animal recipients, typically vertebrate recipients, andpreferably human recipients. A “subject” is a vertebrate, preferably amammal, and most preferably a human. Mammals include, but are notlimited to, humans, farm animals, sport animals, and pets. One of skillin the art can readily vary the parameters of the methods describedherein to accommodate hosts or subjects of variable size and species,including but not limited to, humans of any age.

Advantages of this invention over other methods of tissue engineeringinclude (a) the capability for producing all of the high resolutionthree-dimensional structures required for complex tissues and vitalorgans, and (b) the ability of the mechanical (optionally biodegradable)mold or polymer scaffold to provide support for cell growth and tissueformation, rather than reliance upon biochemical factors alone.

As used herein, the terms “comprises”, “comprising”, and the like canhave the meaning ascribed to them in U.S. Patent Law and can mean“includes”, “including” and the like.

Manufacture of Molds and Polymer Scaffolds

For purposes of this invention a “mold” is a device on the surface ofwhich the branching structure of the microchannels is etched or formed.Fabrication of a mold begins by selection of an appropriate substrate.The choice of a substrate material is guided by many considerations,including the requirements placed on the fabrication process by thedesired mold dimensions, the desired size of the ultimate template, andthe surface properties of the wafer and their interaction with thevarious cell types, extracellular matrix (“ECM”) and polymeric backbone.Also important are the thermal properties, such as the glass transitiontemperature (Tg), which must be high enough so that the network of poresin the mold does not collapse upon solvent removal.

Molds of the present invention can comprise a variety of materials,including, but not limited to, inert materials such as silicon, polymerssuch as polyethylene vinyl acetate, polycarbonate, and polypropylene,and materials such as a ceramic or material such as hydroxyapatite. Inparticular, the mold can comprise from metals, ceramics, semiconductors,organics, polymers, and composites. Representative metals andsemiconductors include pharmaceutical grade stainless steel, gold,titanium, nickel, iron, gold, tin, chromium, copper, alloys of these orother metals, silicon, silicon dioxide. These materials are eitherinherently suitable for the attachment and culture of animal cells orcan be made suitable by coating with materials described herein toenhance cell attachment and culture (e.g. gelatin, matrigel, vitrogenand other tissue culture coatings known in the art).

In an alternative embodiment, MEMS replica molding can be used to make a“polymer scaffold” for seeding cells. In this method, a mold is made asdescribed herein, preferably of silicon (FIG. 1A), and is then used as atemplate on which a polymeric material is cast (FIG. 2). Optionally, thepolymer scaffold can then be peeled away from the mold (FIG. 1B) andseeded with cells.

A “tissue-defining surface” is the surface of a mold or a polymerscaffold, and a “substrate” is the mold or polymer scaffold itself.

The term “polymer” includes polymers and monomers that can bepolymerized or adhered to form an integral unit. The polymer can benon-biodegradable or biodegradable, typically via hydrolysis orenzymatic cleavage. Biodegradable matrices are not typically preferredto construct molds, since they are not implanted and are preferablyreusable. For implantation, polymer scaffolds are preferably used, morepreferably biodegradable polymer scaffolds.

In a preferred embodiment, the biodegradable polymer scaffold comprisesbiodegradable elastomers formed from hydrolyzable monomers as describedin Wang et al., Nature Biotech 20, 602 (2002), the contents of which areincorporated herein by reference. These biodegradable elastomers areanalogous to vulcanized rubber in that crosslinks in a three dimensionalnetwork of random coils are formed. These biodegradable elsatomers arehydrolyzed over time, preferably within 60 days.

Polymer material for implantation should be selected forbiocompatibility. Any degradation products should also be biocompatible.Relatively high rigidity is advantageous so that the polymer scaffoldcan withstand the contractile forces exerted by cells growing within themold. A biocompatible degradable polymer and its degradation productsare non-toxic toward the recipient.

The term “biodegradable” refers to materials that are bioresorbableand/or degrade and/or break down by mechanical degradation uponinteraction with a physiological environment into components that aremetabolizable or excretable, over a period of time from minutes to threeyears, preferably less than one year, while maintaining the requisitestructural integrity. As used in reference to polymers, the term“degrade” refers to cleavage of the polymer chain, such that themolecular weight stays approximately constant at the oligomer level andparticles of polymer remain following degradation. The term “completelydegrade” refers to cleavage of the polymer at the molecular level suchthat there is essentially complete loss of mass. The term “degrade” asused herein includes “completely degrade” unless otherwise indicated.

Materials suitable for polymer scaffold fabrication include, but are notlimited to, poly-dimethyl-siloxane (PDMS), poly-glycerol-sebacate (PGS),polylactic acid (PLA), poly-L-lactic acid (PLLA), poly-D-lactic acid(PDLA), polyglycolide, polyglycolic acid (PGA), polylactide-co-glycolide(PLGA), polydioxanone, polygluconate, polylactic acid-polyethylene oxidecopolymers, modified cellulose, collagen, polyhydroxybutyrate,polyhydroxpriopionic acid, polyphosphoester, poly(αhydroxy acid),polycaprolactone, polycarbonates, polyatnides, polyanhydrides, polyaminoacids, polyorthoesters, polyacetals, polycyanoacrylates, degradableurethanes, aliphatic polyesterspolyacrylates, polymethacrylate, acylsubstituted cellulose acetates, non-degradable polyurethanes,polystyrenes, polyvinyl chloride, polyvinyl fluoride, polyvinylimidazole, chlorosulphonated polyolifins, polyethylene oxide, polyvinylalcohol, teflon RTM, nylon silicon, and shape memory materials, such aspoly(styrene-block-butadiene), polynorbornene, hydrogels, metallicalloys, and oligo(ε-caprolactone)diol as switchingsegment/oligo(p-dioxyanone)diol as physical crosslink. Other suitablepolymers can be obtained by reference to The Polymer Handbook, 3rdedition (Wiley, N.Y., 1989). Combinations of these polymers may also beused.

Polylactide-co-glycolides (PLGA), as well as polylactides (PLA) andpolyglycolides (PGA) have been used to make biodegradable implants fordrug delivery. See U.S. Pat. No. 6,183,781 and references cited therein.Biodegradable materials have been developed for use as implantableprostheses, as pastes, and as templates around which the body canregenerate various types of tissue. Polymers that are both biocompatibleand resorbable in vivo are known in the art as alternatives to autogenicor allogenic substitutes. In a preferred embodiment, polymers areselected based on the ability of the polymer to elicit the appropriatebiological response from cells, for example, attachment, migration,proliferation and gene expression.

Solvents for most of the thermoplastic polymers are known, for example,methylene chloride or other organic solvents Organic and aqueoussolvents for protein and polysaccharide polymers are also known. Thebinder can be the same material as is used in conventional powderprocessing methods or can be designed to ultimately yield the samebinder through chemical or physical changes that occur as a result ofheating, photopolymerization, or catalysis.

Properties of the mold and/or polymer scaffold surface can bemanipulated through the inclusion of materials on the mold or in polymerscaffold material which alter cell attachment (for example, by alteringthe surface charge or structure), porosity, flexibility or rigidity(which may be desirable to facilitate removal of tissue constructs).Moreover, advances in polymer chemistry can aid in the mechanical tasksof lifting and folding as well as the biologic tasks of adhesion andgene expression.

For example, molds can be coated with a unique temperature-responsivepolymer, poly-N-isopropyl acrylamide (PNIPAAm), which demonstrates afully expanded chain conformation below 32° C. and a collapsed, compactconfirmation at high temperatures. When grafted onto surfaces of siliconwafers using electron beam irradiation, it can be used as a temperatureswitch for creating hydrophilic surfaces below 32° C. and hydrophobicsurfaces above 32° C. Since PNIPAAm is insoluble in water over the lowercritical solution temperature (LCST about 32° C.) and reversiblysolubilized below the LCST, cells detach from the substratum by simplylowering the temperature below the LCST. One of skill in the art can (1)engraft the polymer on silicon wafers that are pre-coated withpolystyrene or (2) engraft the polymer on silicon wafers whose surfaceis first modified by vinyl-tricholorosilane. Either of these techniqueswill ensure that the polymer is better integrated and conjugated to itssubstratum (polystyrene in the former case and vinyl groups in the latercase) so that it can serve as an effective thermal switch, useful inreversing cell attachment and detachment as a single contiguous layer ofcells without the usual cell damage.

Another system for promoting both cellular adhesion and lifting of cellsas intact sheets can involve the use of RGD (Arg-Gly-Asp) peptides. TheRGD sequence is part of the domain within the fibronectin molecule thatendows it with the ability to interact with adhesion molecules presenton the cell surface of fibroblasts. Fibronectin itself is awell-characterized extracellular, structural glycoprotein whichinteracts strongly with other extracellular matrix molecules and whichcauses the attachment and spreading of most cells. This function of thefibronectin molecule is localized primarily to the RGD sequence. One ofskill in the art can synthesize RGD peptides with a structural backboneof PMMA that has an RGD peptide sequence at its tips, bound one anotherwith the intermediate layering of polyethylene oxide. This allowsdifferential cell adhesion in only selected areas and not others. Oncethe tissue of desired quality is formed, release of this intactmonolayer of tissue from its substratum is straightforward; it requiresonly the addition of soluble RGD to the culture medium to act as acompetitive substrate to the insolubilized RGD substrate on the siliconmold surface.

In some embodiments, attachment of the cells to the mold and/or polymerscaffold is enhanced by coating the substrate with compounds such asbasement membrane components, agar, agarose, gelatin, gum arabic, typesI, II, III, IV, and V collagen, fibronectin, laminin,glycosaminoglycans, matrigel, vitrogen, mixtures thereof, and othermaterials known to those skilled in the art of cell culture.

Thus, by the methods of the invention, cells can be grown on molds thatare uncoated or coated as described herein, depending upon the materialused for mold construction. Alternatively, cells can be grown on polymerscaffolds made by replica molding techniques.

Micromachining, and Chemical Processing of Silicon and Other MoldMaterials

Molds can be made by creating small mechanical structures in silicon,metal, polymer, and other materials using microfabrication processes.(FIG. 3.) These microfabrication processes are based on well-establishedmethods used to make integrated circuits and other microelectronicdevices, augmented by additional methods developed by workers in thefield of micromachining.

Microfabrication processes that can be used in making the moldsdisclosed herein include lithography; etching techniques, such aslasers, plasma etching, photolithography, or chemical etching such aswet chemical, dry, and photoresist removal; or by solid free formtechniques, including three-dimensional printing (3DP),stereolithography (SLA), selective laser sintering (SLS), ballisticparticle manufacturing (BPM) and fusion deposition modeling (FDM); bymicromachining; thermal oxidation of silicon; electroplating andelectroless plating; diffusion processes, such as boron, phosphorus,arsenic, and antimony diffusion; ion implantation; film deposition, suchas evaporation (filament, electron beam, flash, and shadowing and stepcoverage), sputtering, chemical vapor deposition (CVD), epitaxy (vaporphase, liquid phase, and molecular beam), electroplating, screenprinting, lamination or by combinations thereof. See Jaeger,Introduction to Microelectronic Fabrication (Addison-Wesley PublishingCo., Reading Mass. 1988); Runyan, et al, Semiconductor IntegratedCircuit Processing Technology (Addison-Wesley Publishing Co., ReadingMass. 1990); Proceedings of the IEEE Micro Electro Mechanical SystemsConference 1987-1998; Rai-Choudhury, ed., Handbook of Microlithography,Micromachining & Microfabrication (SPIE Optical Engineering Press,Bellingham, Wash. 1997). The selection of the material that is used asthe mold determines how the surface is configured to form the branchingstructure. The following methods are preferred for making molds.

Typically, micromachining is performed on standard bulk single crystalsilicon wafers of a diameter ranging between about 50 and 300millimeters (mm), preferably approximately 60 mm, and of thicknessranging between about 200 and 1200 μm. These wafers can be obtained froma large number of vendors of standard semiconductor material, and aresawn and polished to provide precise dimensions, uniformcrystallographic orientation, and highly polished, optically flatsurfaces. Wafers made from pyrex borosilicate or other glasses can alsobe procured and inserted into micromachining processes, with alternativeprocesses used to etch the glassy materials.

The geometry of the mold, in particular the number of different featuredepths required, is the major factor determining the specific processsequence. The simplest case is that of a single depth dimension for themold. Specifically, for a silicon substrate, the process sequence (shownin FIG. 3) is as follows: first, the silicon wafer is cleaned, and alayer of photosensitive material is applied to the surface. Typically,the layer is spun on at a high revolution rate to obtain a coating ofuniform thickness. The photoresist is baked, and the wafer is thenexposed to ultraviolet or other short-wavelength light though asemi-transparent mask. This step can be accomplished using any one ofseveral masking techniques, depending on the desired image resolution.The resist is then developed in an appropriate developer chemistry, andthe wafer is then hard-baked to remove excess solvent from the resist.Once the lithographic process has been completed, the wafer can beetched in a plasma reactor using one of several possible chemistries.Etching serves to transfer the two-dimensional pattern into the thirddimension: a specified depth into the wafer. Plasma parameters aredetermined by the desired shape of the resulting trench (semi-circular,straight-walled profile, angled sidewall), as well as by the selectivityof the etchant for silicon over the masking photoresist. Once theetching has been completed, the photoresist can be removed and the waferprepared for use in the tissue molding, process.

Increased flexibility in the geometry of wafer mold can be obtained byinserting additional cycles of masking and etching, as shown in FIG. 4.Here, a second step in which a masking layer has been applied, and openareas etched, is shown. This modification provides the opportunity tomachine channels of varying depths into the wafer mold. To design a moldthat is suitable for the culturing of endothelial cells, increasedflexibility is very important due to the need for vascular branches withdifferent diameters. The techniques can be extended to provide as manyadditional layers and different depths as are desired. In addition,these techniques can be used to create secondary patterns within thepattern of microchannels. For example, it may be advantageous to havewells within the microchannels for culturing additional cell types suchas feeder cells. The pattern of microchannels also can be designed tocontrol cell growth, for example, to selectively control thedifferentiation of cells.

Glass and polymeric wafer molds can be fabricated using a similarsequence, but the actual process can be modified by the addition of anintervening masking layer, since etchants for these materials may attackphotoresist as well. Such intervening materials simply function totransfer the pattern from the photoresist to interlayer and then on tothe wafer below. For silicon etched in one of several wet chemistries,an intervening layer may also be necessary.

Electrolytic anodization of silicon in aqueous hydrofluoric acid,potentially in combination with light, can be used to etch channels intothe silicon. By varying the doping concentration of the silicon wafer tobe etched, the electrolytic potential during etching, the incident lightintensity, and the electrolyte concentration, control over the ultimatepore structure can be achieved. This process uses deep plasma etching ofsilicon. Needles are patterned directly using photolithography, ratherthan indirectly by controlling the voltage (as in electrochemicaletching), thus providing greater control over the final mold geometry.

In this process, an appropriate masking material (e.g., metal) isdeposited onto a silicon wafer substrate and patterned into dots havingthe diameter of the desired channels. The wafer is then subjected to acarefully controlled plasma based on fluorine/oxygen chemistries to etchvery deep, high aspect ratio trenches into the silicon. See, e.g.,Jansen, et al., “The Black Silicon Method IV: The Fabrication ofThree-Dimensional Structures in Silicon with High Aspect Ratios forScanning Probe Microscopy and Other Applications,” IEEE Proceedings ofMicro Electro Mechanical Systems Conference, pp. 88-93 (1995).

A metal layer is first evaporated onto a planar substrate. A layer ofphotoresist is then deposited onto the metal to form a patterned mold,which leaves an exposed-metal region in the shape of needles. Byelectroplating onto the exposed regions of the metal seed layer, themold bounded by photoresist can be filled with electroplated material.Finally, the substrate and photoresist mold are removed, leaving thefinished mold array. The molds produced by this process generally havechannels with diameters on the order of about 1 μm or larger.Preferably, microchannels have a diameter of about 1 μm to about 500 μm.See Frazier, et al., “Two dimensional metallic microelectrode arrays forextracellular stimulation and recording of neurons”, IEEE Proceedings ofthe Micro Electro Mechanical Systems Conference, pp. 195-200 (1993).

Another method of forming solid silicon molds is by using epitaxialgrowth on silicon substrates, as is utilized by Containerless Research,Inc. (Evanston, Ill., USA) for its products.

The size distribution of the etched porous structure is highly dependenton several variables, including doping kind and illumination conditions,as detailed in Lehmann, “Porous Silicon—A New Material for MEMS”, IEEEProceedings of the Micro Electro Mechanical Systems Conference, pp. 1-6(1096). Porous polymer molds can be formed, for example, by micromoldinga polymer containing a volatilizable or leachable material, such as avolatile salt, dispersed in the polymer, and then volatilizing orleaching the dispersed material, leaving a porous polymer matrix in theshape of the mold. Hollow molds can be fabricated, for example, usingcombinations of dry etching processes (Laermer, et al., “Bosch DeepSilicon Etching: Improving Uniformity and Etch Rate for Advanced MEMSApplications,” Micro Electro Mechanical Systems, Orlando, Fla., USA,(Jan. 17-21, 1999); Despont, et al., “High-Aspect-Ratio, Ultrathick,Negative-Tone Near-UV Photoresist for MEMS”, Proc. of IEEE 10^(th)Annual International Workshop on MEMS, Nagoya, Japan, pp. 518-522 (Jan.26-30, 1997)); micromold creation in lithographically-defined polymersand selective sidewall electroplating; or direct micromolding techniquesusing epoxy mold transfers.

A chromium mask can be substituted for the solid molds using a siliconnitride layer covered with chromium. Solid molds are then etched, thechromium is stripped, and the silicon is oxidized. The silicon nitridelayer will prevent oxidation. The silicon nitride is then stripped,leaving exposed silicon and oxide-covered silicon everywhere else. Theneedle is then exposed to an ICP plasma which selectively etches thesilicon in a highly anisotropic manner to form the interior hole of theneedle. A second method uses solid silicon as ‘faints’ around which theactual needle structures are deposited. After deposition, the forms areetched away, yielding the hollow structures. Silica needles or metalneedles can be formed using different methods. The wafers are thenoxidized to a controlled thickness, the silicon nitride is then strippedand the silicon core selectively etched away (e.g., in a wet alkalinesolution) to form a hollow silica mold.

In another embodiment, deep reactive ion etching is combined with amodified black silicon process in a conventional reactive ion etcher.First, designs are patterned through photoresist into SiO₂, such as on asilicon wafer. Then the silicon can be etched using deep reactive ionetching (DRIE) in an inductively coupled plasma (ICP) reactor to etchdeep vertical holes or channels. The photoresist is then removed. Next,a second photolithography step patterns the remaining SiO₂ layer. Thephotoresist is then removed and the silicon wafer again deep siliconetched completely through the wafer in the regions not covered withSiO₂). (See FIG. 5.) This process can be varied as follows. After thewafer is patterned, the photoresist and SiO₂ layers are replaced withconformal DC sputtered chromium. The second JCP etch is replaced with aSF₆/O₂ plasma etch in a reactive ion etcher (RIE), which results inpositively sloping outer sidewalls. Henry, et al., “MicromachinedNeedles for the Transdermal Delivery of Drugs,” Micro Electra MechanicalSystems, Heidelberg, Germany, pp. 494-498 (Jan. 26-29, 1998).Alternatively, silicon may be etched anisotropically using Deep ReactiveIon Etching (DRIE) using a switched-process technology (A. A. Ayon, S.Nagle, L. Frechette, A. Epstein and M. A. Schmidt, “Tailoring etchdirectionality in a deep reactive ion etching tool,” J. Vac. Sci. Tech.B 18, 1412 (2000)).

Metal shapes can be formed by physical vapor deposition of appropriatemetal layers on solid forms, which can be made of silicon using thetechniques described above, or which can be formed using other standardmold techniques such as embossing or injection molding. The metals areselectively removed using electropolishing techniques, in which anapplied anodic potential in an electrolytic solution will causedissolution of metals due to concentration of electric field lines. Oncethe underlying silicon forms have been exposed, the silicon isselectively etched away to form structures. This process could also beused to make structures made from other materials by depositing amaterial other than metal on the needle forms and following theprocedure described above.

Molds formed of silicon dioxide can be made by oxidizing the surface ofthe silicon mold forms, rather than depositing a metal and then etchingaway the solid needle forms to leave the hollow silicon dioxidestructures. In one embodiment, hollow, porous, or solid molds areprovided with longitudinal grooves or other modifications to theexterior surface of the molds.

Polymeric molds can also be made using microfabrication. For example,the epoxy molds can be made as described above, and injection moldingtechniques can be applied to form the structures. Thesemicromicromolding techniques are relatively less expensive to replicatethan the other methods described herein.

Three dimensional printing (3DP) is described by Sachs, et al.,Manufacturing Review 5, 117-126 (1992) and U.S. Pat. No. 5,204,055 toSachs, et al. 3DP is used to create a solid object by ink jet printing abinder into selected areas of sequentially deposited layers of powder.Each layer is created by spreading a thin layer of powder over thesurface of a powder bed. The powder bed is supported by a piston, whichdescends upon powder spreading and printing of each layer (or,conversely, the ink jets and spreader are raised after printing of eachlayer and the bed remains stationary). Instructions for each layer arederived directly from a computer-aided design (CAD) representation ofthe component. The area to be printed is obtained by computing the areaof intersection between the desired plane and the CAD representation ofthe object. The individual sliced segments or layers are joined to formthe three-dimensional structure. The unbound powder supports temporarilyunconnected portions of the component as the structure is built but isremoved after completion of printing.

SPF methods other than 3DP that can be utilized to some degree asdescribed herein are stereo-lithography (SLA), selective laser sintering(SLS), ballistic particle manufacturing (BPM), and fusion depositionmodeling (FDM). SLA is based on the use of a focused ultraviolet (UV)laser that is vector scanned over the top of a bath of aphotopolymerizable liquid polymer material. The UV laser causes the bathto polymerize where the laser beam strikes the surface of the bath,resulting in the creation of a first solid plastic layer at and justbelow the surface. The solid layer is then lowered into the bath and thelaser generated polymerization process is repeated for the generation ofthe next layer, and so on, until a plurality of superimposed layersforming the desired apparatus is obtained. The most recently createdlayer in each case is always lowered to a position for the creation ofthe next layer slightly below the surface of the liquid bath. A systemfor stereolithography is made and sold by 3D Systems, Inc., of Valencia,Calif., which is readily adaptable for use with biocompatible polymericmaterials. SLS also uses a focused laser beam, but to sinter areas of aloosely compacted plastic powder, the powder being applied layer bylayer. In this method, a thin layer of powder is spread evenly onto aflat surface with a roller mechanism. The powder is then raster-scannedwith a high-power laser beam. The powder material that is struck by thelaser beam is fused, while the other areas of powder remain dissociated.Successive layers of powder are deposited and raster-scanned, one on topof another, until an entire part is complete. Each layer is sintereddeeply enough to bond it to the preceding layer. A suitable systemadaptable for use in making medical devices is available from DTMCorporation of Austin, Tex.

BPM uses an ink-jet printing apparatus wherein an ink-jet stream ofliquid polymer or polymer composite material is used to createthree-dimensional objects under computer control, similar to the way anink: jet printer produces two-dimensional graphic printing. The mold isformed by printing successive cross-sections, one layer after another,to a target using a cold welding or rapid solidification technique,which causes bonding between the particles and the successive layers.This approach as applied to metal or metal composites has been proposedby Automated Dynamic Corporation of Troy, N.Y. FDM employs an x-yplotter with a z motion to position an extrudable filament formed of apolymeric material, rendered fluid by heat or the presence of a solvent.A suitable system is available from Stratasys, Incorporated ofMinneapolis, Minn.

The design of the channels in the mold can be constructed by a number ofmeans, such as fractal mathematics, which can be converted by computersinto two-dimensional arrays of branches and then etched onto wafers.Also, computers can model from live or preserved organ or tissuespecimens three-dimensional vascular channels, convert totwo-dimensional patterns and then help in the reconversion to athree-dimensional living vascularized structure. Techniques forproducing the molds include techniques for fabrication of computer chipsand microfabrication technologies. Other technologies include lasertechniques.

Design of Apparatus

As shown in FIG. 6, in a preferred embodiment, the pattern in the mold(10), formed of a silicon wafer (11), begins with one or more largechannels (12), which serially branch into a large array of channels assmall as individual capillaries (14 a, 14 b, 14 c, etc.), then convergeto one or more large channels (16). The cross-section of the single“arterial” channel (12) and “venous” channel (16) is shown in FIG. 7A.The cross-section of the portion of mold (10) containing the “capillary”channels (14 a, 14 b, 14 c, etc.), is shown in FIG. 7A. The mold isshown in cross-section in FIG. 7C, with a depth of approximately 5 μm.

The etched surface serves as a template for the circulation of anindividual tissue or organ. Living endothelial cells seeded into thesechannels and provided with flow of appropriate nutrients and gases willline the channels to form blood vessels. In one embodiment, as shown inFIG. 8A, mold and/or polymer scaffold pieces (30 and 32) are fittedtogether to make an enclosure (34), and the cells are cultured. Thevascular cells form vascular channels (36) based on the pattern etchedin the mold, as shown in FIG. 8B. In Example 1, it has been demonstratedthat cells seeded onto surfaces of silicon and pyrex will lay downmatrix and form sheets of tissue of the cell type of origin, eitherhepatic or endothelial. Once formed and sustained by their own matrix,the top of the mold or polymer scaffold (32) can be removed, and theorgan or tissue specific cells can then be added to the etched surface,where they attach and proliferate to form a thin, vaseularized sheet oftissue (36). As shown in FIG. 8C, the tissue can then be gently liftedfrom the mold or polymer scaffold using techniques such as fluid flowand other supporting material, as necessary. Alternatively, the polymercan be degraded. These sheets can then be formed into three-dimensionalunits of tissue. In effect, the wafer of silicon or pyrex or the polymerscaffold has acted as a template for the formation of tissue.

In a more preferred embodiment, as shown in FIG. 9A, mold and/or polymerscaffold pieces (1 and 3) are fitted together and separated by asemi-permeable membrane (2). The vascular cells are seeded into onelayer and cultured to form vascular channels (4) based on the patternetched in the surface of the mold, as shown in FIG. 9A. The organ ortissue specific cells are added to the second patterned surface, wherethey attach and proliferate (5) to form a vaseularized tissue bilayer.The second patterned surface optionally comprises inlets for optionallyincludes inlets for neural inervation, urine flow, biliary excretion orother activity.

Semi-Permeable Membrane

A semi-permeable membrane can be used to separate the first mold orpolymer scaffold from the second mold or polymer scaffold in themicrofabricated apparatuses of the invention. Preferably, the pore sizeof the membrane is smaller than the cell diameters, thus, cells will notbe able to pass through (i.e. a low permeability for animal cells),while low molecular weight nutrients and fluids can pass through (i.e. ahigh permeability for nutrients), thereby providing adequatecell-to-cell signaling. Cell sizes vary but in general, they are in therange of microns. For example, a red blood cell has a diameter of 8 μm.Preferably, the average membrane pore size is on a submicron-scale toensure effective screening of the cells.

Semi-permeable membranes of the present invention comprise a wide arrayof different membrane types and morphologies, which can be classified asfollows:

(1) Track-etch membranes consisting of cylindrical through-holes in adense polymer matrix. These membranes are typically made by ion-etching;or

(2) Fibrous membranes made by various deposition techniques of polymericfibers. While these membranes do not have a well-defined pore topology,production methods have been sufficiently refined so that fibrousmembranes have specific molecular weight cut-offs.

Track-etch type membranes are preferred, as they limit the fluid motionin one direction. Preferably, fluid motion is in the vertical direction.Fibrous membranes permit fluid motion both laterally and vertically.

The development of an appropriate membrane will mirror the deviceprogression. Biocompatible and non-degradable membranes can beincorporated in microchannels that are made from poly(dimethyl siloxane)(PDMS). Since PDMS is non-degradable, the membranes do not need to bedegradable either. However, degradable membranes and materials formicrochannels can also be used. There exists a variety of commercialtrack-etch membranes with well-defined pore sizes that can be used forthis purpose. Care must be taken to properly incorporate the membranesinto the existing microchannels without leaking. To this end, themembranes can be bonded with either an oxygen plasma or a silicone-basedadhesive. A small recession can be designed into the microchannels sothat the membrane can fit tightly therein.

In principle, membrane formation from polymers relies on phase-phaseseparation. Polymer-solvent interactions are complex, and polymer phasediagrams are significantly more complicated than those for monomericmaterials, e.g., metals. Phase separation can be induced either bydiffusion (diffusion-induced phase separation or “DIPS”) or by thermalmeans (thermal induced phase separation or “TIPS”).

A DIPS system comprises polymer, solvent and non-solvent. The polymersolution is cast as a thin film and then immersed in a coagulation bathcontaining the non-solvent. This process is governed by the diffusion ofvarious low molecular weight components. The exchange of solvent andnon-solvent between the polymer solution and the coagulation bath leadsto a change in the composition in the film and phase separation isinduced. After some time, the composition of the polymer-rich phasereaches the glass transition composition and the system solidifies. Toavoid macrovoid formation, a small amount of non-solvent can be mixedwith the polymer solution. In a preferred embodiment, the polymer ispolycaprolactone (PCL) and the separation system is chloroform/methanol.Specifically, a polymer solution with a concentration ranging from about5-10% wt. is made. PCL is prepared by dissolving it in chloroform atroom temperature under gentle stirring. Once the polymer has completelydissolved, a small amount is placed on a clean mirror surface, and amembrane knife is used to spread out a film with preset thickness. Thethickness of the film can be adjusted by changing the gap between theknife blade and the mirror surface. Once the film has been spread, theentire mirror is immersed in a methanol bath. Phase separation occursalmost instantaneously, but the film and mirror are left in thecoagulation bath for up to about 10 minutes to lock in the morphology. Atypical membrane thickness is about 100 μm, and the pore size is on theorder of about 1 μm, preferably between about 0.01 and 20 μm (FIG. 10).Membrane morphology can be varied by altering thecomposition/concentration of the polymer solution, the film thickness,the components of the coagulation bath, and/or the process conditions.One skilled in the art would understand how to vary any one of theparameters to achieve the desired result.

A TIPS system comprises a thermal gradient to induce phase separation.By choosing a polymer-solvent system that is miscible at hightemperatures, but immiscible at low temperatures, e.g., roomtemperature, phase separation can be induced upon cooling down thepolymer solution. In a preferred embodiment, the polymer is PCL and theseparation system is DMF/10% C₃H₈O₃.

Cells to be Seeded onto the Mold or Polymer Scaffold

The tissue will typically include one or more types of functional,mesenchymal or parenchymal cells, such as smooth or skeletal musclecells, myocytes (muscle stem cells), fibroblasts, chondrocytes,adipocytes, fibromyoblasts, ectodermal cells, including ductile and skincells, hepatocytes, kidney cells, pancreatic islet cells, cells presentin the intestine, and other parenchymal cells, osteoblasts and othercells forming bone or cartilage, and hematopoictic cells. In some casesit may also be desirable to include nerve cells, The vasculature willtypically be formed from endothelial cells. “Parenchymal cells” includethe functional elements of an organ, as distinguished from the frameworkor stroma. “Mesenchymal cells” include connective and supportingtissues, smooth muscle, vascular endothelium and blood cells.

Cells can be obtained by biopsy or harvest from a living donor, cellculture, or autopsy, all techniques well known in the art. Cells arepreferably autologous. Cells to be implanted can be dissociated usingstandard techniques such as digestion with a collagenase, trypsin orother protease solution and are then seeded into the mold or polymerscaffold immediately or after being maintained in culture. Cells can benormal or genetically engineered to provide additional or normalfunction. Immunologically inert cells, such as embryonic or fetal cells,stem cells, and cells genetically engineered to avoid the need forimmunosuppression can also be used. Methods and drugs forimmunosuppression are known to those skilled in the art oftransplantation.

Undifferentiated or partially differentiated precursor cells, such asembryonic gel axe cells (Gearhart, et al., U.S. Pat. No. 6,245,566),embryonic stem cells (Thomson, U.S. Pat. Nos. 5,843,780 and 6,200,802),mesenchymal stem cells (Caplan, et al. U.S. Pat. No. 5,486,359), neuralstem cells (Anderson, et al., U.S. Pat. No. 5,849,553), hematopoieticstem cells (Tsukarnoto, U.S. Pat. No. 5,061,620), multipotent adult stemcells (Furcht, et al., WO 01/11011) can be used in this invention. Cellscan be kept in an undifferentiated state by co-culture with a fibroblastfeeder layer (Thomson, U.S. Pat. Nos. 5,843,780 and 6,200,802), or byfeeder-free culture with fibroblast conditioned media (Xu, et al. Nat.Biotechnol., 19, 971 (2001)). Undifferentiated or partiallydifferentiated precursor cells can be induced down a particulardevelopmental pathway by culture in medium containing growth factors orother cell-type specific induction factors or agents known in the art.Some examples of such factors are shown in Table 1.

TABLE 1 Selected Examples of Differentiation Inducing Agents AgentProgenitor Differentiated Cell Vascular Endothelial Growth EmbryonicStem Cell Hematopoietic Cell¹ Factor Sonic Hedgehog Floor Plate MotorNeuron² Insulin-like Growth Factor II Embryonic Stem Cell Myoblast³Osteogenin Osteoprogenitor Osteoblast⁴ Cytotoxic T Cell DifferentiationSpleen Cell Cytotoxic T Lymyphocyte⁵ Factor β-catenin Skin Stem CellFollicular Keratinocyte⁶ Bone Morphogenic Protein 2 Mesenchymal StemCell Adipocytes, Osteoblasts⁷ Interleukin 2 Bone Marrow PrecursorNatural Killer Cells⁸ Transforming Growth Factor β Cardiac FibroblastCardiac Myocyte⁹ Nerve Growth Factor Chromaffin Cell SympatheticNeuron¹⁰ Steel Factor Neural Crest Melanocyte¹¹ Interleukin 1Mesencephalic Progenitor Dopaminergic Neuron¹² Fibroblast Growth Factor2 GHFT Lactotrope¹³ Retinoic Acid Promyelocytic Leukemia Granulocyte¹⁴Wnt3 Embryonic Stem Cell Hematopoietic Cell¹⁵ ¹Keller, et al. (1999)Exp. Hematal 27: 777-787. ²Marti, et al. (1995) Nature. 375: 322-325.³Prelle, et al. (2000) Biochem. Biophy. Res. Commun. 277: 631-638.⁴Amedee, et al. (1994) Differentiation. 58: 157-164. ⁵Hardt, et al.(1985) Eur. J. Immunol. 15: 472-478. ⁶Huelsken et al. (2001) Cell. 105:533-545. ⁷Ji, et al. (2000) J. Bone Miner. Metab. 18: 132-139.⁸Migliorati, et al. (1987) J. Immunol. 138: 3618-3625. ⁹Eghbali, et al.(1991) Prob. Natl. Acad. Sci. USA. 88: 795-799. ¹⁰Niijima, et al. (1995)J. Neurosci. 15: 1180-1194. ¹¹Guo, et al. (1997) Dev. Biol. 184: 61-69.¹²Ling, et al. (1998) Exp. Neurol. 149: 411-423. ¹³Lopez-Fernandez, etal. (2000) J. Biol. Chem. 275: 21653-60. ¹⁴Wang, et al. (1989) Leuk.Res. 13: 1091-1097. ¹⁵Lako, et al. (2001) Mech. Dev. 103: 49-59.

A stem cell can be any known in the art, including, but not limited to,embryonic stem cells, adult stem cells, neural stem cells, muscle stemcells, hematopoietic stem cells, mesenchymal stem cells, peripheralblood stem cells and cardiac stem cells. Preferably, the stem cell ishuman. A “stem cell” is a pluripotent, multipotent or totipotent cellthat can undergo self-renewing cell division to give rise tophenotypically and genotypically identical daughter cells for anindefinite time and can ultimately differentiate into at least one finalcell type.

The quintessential stem cell is the embryonal stem cell (ES), as it hasunlimited self-renewal and multipotent and or pluripotentdifferentiation potential, thus possessing the capability of developinginto any organ, tissue type or cell type. These cells can be derivedfrom the inner cell mass of the blastocyst, or can be derived from theprimordial germ cells from a post-implantation embryo (embryonal germcells or EG cells). ES and EG cells have been derived from mice, andmore recently also from non-human primates and humans. Evans et al.(1981) Nature 292:154-156; Matsui et al. (1991) Nature 353:750-2;Thomson et al. (1995) Proc. Natl. Acad. Sci. USA. 92:7844-8; Thomson etal. (1998) Science 282:1145-1147; and Shamblott et al. (1998) Proc.Natl. Acad. Sci. USA 95:13726-31.

The terms “stem cells,” “embryonic stem cells,” “adult stem cells,”“progenitor cells” and “progenitor cell populations” are to beunderstood as meaning in accordance with the present invention cellsthat can be derived from any source of adult tissue or organ and canreplicate as undifferentiated or lineage committed cells and have thepotential to differentiate into at least one, preferably multiple, celllineages.

The hepatocytes added to the apparatus of the invention can be highlyproliferative hepatocytes, known as small hepatocytes (SHCs), which havethe ability to proliferate in vitro for long periods of time (Mitaka, etal., Biochem Biophys Res Commun 214, 310 (1995); Taneto, et al, Am JPathol 148, 383 (1996)). Small hepatocytes express hepatocyte specificfunctions such as albumin production (Mitalca, et al., Hepatology 29,111 (1999)). The survival and function of small hepatocytes whenco-cultured on three-dimensional micronorous biodegradable polymer moldsunder dynamic culture conditions was demonstrated (Example 4).

Methods for Seeding Cells into Molds or Polymer Scaffolds

Cell Seeding

After the mold with the desired high degree of micromachining isprepared, the molds themselves or polymer scaffolds are seeded with thedesired cells or sets of cells. The distribution of cells throughout themold or polymer scaffold can influence both (1) the development of avascularized network, and (2) the successful integration of the vasculardevice with the host. The approach used in this invention is to providea mechanism for the ordered distribution of cells onto the mold orpolymer scaffold. Cells that are enriched for extracellular matrixmolecules or for peptides that enhance cell adhesion can be used. Cellscan be seeded onto the mold or polymer scaffold in an ordered mannerusing methods known in the art, for example, Teebken, et al., Eur J.Vasa Endovasc. Surg. 19, 381 (2000); Ranucci, et al., Biomaterials 21,783 (2000). Also, tissue-engineered devices can be improved by seedingcells throughout the polymeric scaffolds and allowing the cells toproliferate in vitro for a predetermined amount of time beforeimplantation, using the methods of Burg et al., J. Biomed. Mater. Res51, 642 (2000).

For purposes of this invention, “animal cells” can comprise endothelialcells, parenchymal cells, bone marrow cells, hematopoietic cells, musclecells, osteoblasts, stem cells, mesenchymal cells, stem cells, embryonicstem cells, or fibroblasts. Parenchymal cells can be derived from anyorgan, including heart, liver, pancreas, intestine, brain, kidney,reproductive tissue, lung, muscle, bone marrow or stem cells.

In one embodiment, the mold or polymer scaffold is first seeded with alayer of parenchymal cells, such as hepatocytes or proximal tubulecells. This layer can be maintained in culture for a week or so in orderto obtain a population doubling. It can be maintained in a perfusionbioreactor to ensure adequate oxygen supply to the cells in theinterior. The apparatus is then seeded with a layer of endothelial cellsand cultured further. In regions where the matrix is resorbed rapidly,the tissue can expand and become permeated with capillaries.

Cell Seeding of Horizontal Layer by Laminar Flow.

A structure comprising joined or fastened molds and/or polymerscaffolds, with or without a semi-permeable membrane between them, iscalled an “apparatus” for purposes of this invention. Sets of cells canbe added to or seeded into the three-dimensional apparatuses, which canserve as a template for cell adhesion and growth by the added or seededcells. The added or seeded cells can be parenchymal cells, such ashepatocytes or proximal tubule cells. Stem cells can also be used. Asecond set of cells, such as endothelial cells, can be added to orseeded onto the assembled apparatus through other vessels than thoseused to seed the first set of cells. The cell seeding is performed byslow flow. As a practical matter, the geometry of the apparatus willdetermine the flow rates. In general, endothelial cells can enter andform vessel walls in micromachined channels that are about 10-50 μm.Thus, in addition to serving as a mechanical framework for the organ,the assembled apparatus provides a template for all of themicrostructural complexity of the organ, so that cells have a mechanicalmap to locate themselves and form subsystems, such as blood vessels inthe liver.

Optionally, functional cells are seeded into both a first and secondmold and/or polymer scaffold with microchannels on their surfaces, andthe two molds and/or polymer scaffolds are joined or fastened with asemi-permeable membrane between them, allowing gas exchange, diffusionof nutrients, and waste removal. One layer comprises the circulationthrough which blood, plasma or media with appropriate levels of oxygencan be continuously circulated to nourish the second layer. The secondlayer comprises a reservoir for the functional cells of an organ, andoptionally includes inlets for neural inervation, urine flow, biliaryexcretion or other activity. This results in an apparatus for makingtissue lamina, wherein each of the first and second molds and/or polymerscaffolds and the semi-permeable membrane are comprised of material thatis suitable for attachment and culturing of animal cells. The sheet oftissue created by the apparatuses and/or methods of the invention isreferred to as “tissue lamina”.

Channels in the horizontal direction typically proceed from larger tosmaller to larger. The geometries can be as complex as desired in-plane(horizontal direction). Thus, one can use small geometries in-plane(such as horizontal conduits of about 5-20 μm). The alignment ofthrough-holes creates vertical conduits or channels in the z-axis.However, the vertical channels need not go from larger to smaller tolarger. In the vertical direction, the vertical channels are typicallyparallel to each other and have diameters on the micron level, largeenough only to allow cell seeding (e.g., hepatocytes are about 40 μm).In one embodiment, different types of cells are seeded horizontally ontodifferent layers of the assembled apparatus. In another embodiment, thedifferent types of cells are seeded using pores or channels fromdifferent directions. Various combinations are also possible. (See FIG.11.)

Although described herein with particular reference to formation ofvascularized tissue, it should be understood that the channels can beused to form lumens for passage of a variety of different fluids, notjust blood, but also bile, lymph, nerves, urine, and other body fluids,and for the guided regeneration or growth of other types of cells,especially nerve cells. The tissue layer can include some lumens forforming vasculature and some for other purposes, or be for one purpose,typically providing a blood supply to carry oxygen and nutrients to andfrom the cells in the tissue.

Molecules such as growth factors or hormones can be covalently attachedto the surface of the molds and/or polymer scaffolds and/orsemi-permeable membrane to effect growth, division, differentiation ormaturation of cells cultured thereon.

Construction of Tissue or Organ Equivalents

Engineered tissue lamina can be systematically folded and compacted intoa three-dimensional vascularized structure, as shown in FIGS. 12 and 13.The two-dimensional surface of the mold can be varied to aid in thefolding and compacting process. For example, the surface can be changedfrom planar to folded accordion-like. It can be stacked into multipleconverging plates. It could be curvilinear or have multiple projections.

Different types of tissue, or multiple layers of the same type oftissue, can be superposed prior to folding and compacting, to createmore complex or larger structures. For example, a tubular system can belayered onto a vascular system to fabricate glomerular tissue andcollecting tubules for kidneys. Bile duct tubes can be overlaid onvascularized liver or hepatocyte tissue, to generate a bile ductdrainage system. Alveolar or airway tissue can be placed on lungcapillaries to make new lung tissue. Nerves or lymphatics can be addedusing variations of these same general techniques.

FIGS. 12A-G are perspective views of ways in which a single tissuelamina (FIG. 12A) can be folded (FIGS. 12B and 12C) or stacked (FIG.12D), or expanded to form a balloon shape (FIG. 12E), funnel (FIG. 12F),or large lumen (FIG. 12G).

In addition to embodiments in which a single tissue layer is formed, theapparatus shown in FIG. 9 can be used for three dimensional tissue andorgan formation. The addition of the second mold or polymer scaffoldallows the functional unit of the organ to be added, and likewise allowsprecision for patterning of exocrine outflow. For example, in the liver,the parenchymal cells are hepatocytes and the exocrine system is thebiliary system. By the addition of the second compartment containinghepatocyes and biliary cells, the functional tissue of the liver can beachieved and biliary excretion can be designed and enfolded.

This patterning can be made more complex with the addition of furtherlayers separated by permeable membranes. Several molds and/or polymerscaffolds, with or without semi-permeable membranes between them, can bestacked in rational arrays to produce complex tissue in 3-dimensionalspace (see FIG. 28, for example). These layers of molds and/or polymerscaffolds, and optionally, semi-permeable membranes, can beappropriately interdigitated and connected (e.g. via through-holes) toproduce vascular connections through the depths of the stack, as well asexcretory outflow systems through the depths of the tracts.

Stacking Molds and/or Polymer Scaffolds to Achieve Three Dimensionality.

Extension of the two-dimensional technology into the third dimension canbe accomplished by stacking the two-dimensional layers on top of eachother. This stacking method begins with many molds and/or polymerscaffolds produced by the techniques described in previous sections.Once these molds and/or polymer scaffolds (nominally of the same size)are created, they are lain down or bonded to other separate molds and/orpolymer scaffolds, atop one another. The layers can be connected bythrough-holes, which extend through the z-axis of the molds and/orpolymer scaffolds. The pattern of microchannels on the surface of eachmold or polymer scaffold can differ or be similar to the previous layer,depending upon fluid mechanical considerations. In addition to thetwo-dimensional channels embedded in each layer, the through-holes canprovide vessel structures that extend up into the third (vertical)dimension. Each successive layer could have slightly different patternsof through-holes, on that the effect would be to have vessels extendinginto the third dimension that are not necessarily preciselyperpendicular to the plane of the sheet.

By extending this technology as needed, one can move from the presentlyachievable formation of small (˜100 cm²) tissue sheets, each containingone plane of blood vessels, to the formation of perhaps 100 cm³ ofmaterial, enough to build an organ. The process is low-cost, scalable,can be customized for the physiology of a particular patient, and isbased upon currently available microfabrication technology.

Fastening the Stacked Layers.

An aspect of this invention is the fastening or sealing of the polymericmold layers. Preferably, the layers are irreversibly bound beforeimplantation into the host. Depending on the composition of the layeredmaterial, the layers can be sealed by solvent bonding; reflow by heating(40° C.); treating surface with oxygen plasma; or by polymer flow at thesurface. Biocompatible polymer materials maybe bonded together by plasmaactivation to form sealed structures (Jo et al., SPIE 3877, 222 (1999)).The basic process results in bonded layers with channel architectureclosely resembling that obtained with silicon etched molds.

Silicon-Glass Microfluidic Chambers to Test Sealing of Stacks.

Microfluidic tests have been performed that demonstrate that bondedapparatuses are leakproof and support fluid pressures necessary fordynamic cell seeding. (See Example 3.) One of the most common methodsused to seal micromachined wafers together is anodic bonding, atechnique based on the high concentration of mobile ions in many glasses(Camporese, et al., IEEE Electron. Device Lett. EDL 2, 61(1981)). Thisprocess produces a permanent seal; fracture testing of silicon-glassanodically bonded interfaces produces a failure within the bulk of theglass.

Etched wafers maybe bonded together, producing closed lumens suitablefor fluidic experiments. A fluidic test was performed with a mixed-phaseflow of alcohol with 10 μm fluorescent microspheres. An unetched glasscapping layer was mechanically drilled for inlet and outlet fluid ports,and then anodic ally bonded to a silicon wafer plasma-etched with theTEP-1-geometry. A permanent seal with no leaks was produced, enablingone to obtain highly accurate pressure and flow data.

Alternatively, the multilayer device of the invention can be configuredsuch that each of the layers has an alignment indentation on one surfaceof the layer and an alignment protrusion on the opposing surface ofanother layer. The alignment indentations shaped to mate with thealignment protrusion, so that the layers are held together.

Alternative Methods of Stacking.

To build up the mold and/or polymer scaffold layers by mechanicalassembly, the layers can be mechanically mated using biodegradable ornon-biodegradable barbs, pins, screws, clamps, staples, wires, string,or sutures. (See, U.S. Pat. No. 6,143,293.) With this mechanicalassembly approach, each prefabricated section can comprise differentmold and/or polymer scaffold material and/or different moldmicrostructures. Different sections of these can be seeded with cellsbefore assembly. Cells thus be can be embedded into the mold or polymerscaffold by assembling sections around these components. In addition,surface features on each mold, which are readily fabricated, become partof the internal microstructure (e.g., molded surface channels becomeconduits for cell infusion, or for blood flow to stimulateangiogenesis). A surface feature on an individual mold or polymerscaffold will become an internal feature when another segment isassembled over it. For example, surface features such as channels can bemicromachined into a first mold or polymer scaffold layer. When a secondmold or polymer scaffold layer is placed atop that a first layer, themicromachined surface feature becomes an internal feature of theapparatus.

Rolling or Folding to Achieve Three Dimensionality

An alternate method for achieving three-dimensionality is to generate along strip of polymer mold material, which contains repeating units ofthe blood vessel network along with through-holes, and to fold the moldfilm in a z-fold fashion while aligning the through-holes to oneanother.

The rolling or folding process begins with the generation of a lengthystrip of polymer mold material, which contains a serial array of unitcells each of which is comprised of an array of channels mimicking thevascular network, produced from a wafer mold by molding, embossing, orthe like. These unit cells can be identical or can be different. Theunits are linked to through-holes that provide the vertical channelconnections between horizontal blood vessel layers. Once the polymericscaffold strip has been formed, it is folded in a z-fold fashion (FIG.12C), and bonded together so that each fold is attached to the filmportions above and below it with alignment to the through-holes.

This roll can be of a length to provide sufficient scaffolding materialfor an entire human organ, which can be hundreds or even more multiplesof the area of a single wafer. Each section of the roll is a sheet ofpolymeric mold with closed lumens, or vessels. The vessels in eachfolded section of sheet are connected to a through-hole at the edge ofthe sheet (for example, one on each side, for inlet and outlet bloodflow). During folding, the sheet sections are folded such that thethrough-hole openings align, forming a vessel in the third (z)dimension.

The roll can be in the shape of a spiral, helix, jelly roll or othercylindrically shaped objects. (See FIG. 12.)

The described three-dimensional tissue structures can then be imp/antedinto animals or patients by directly connecting the blood vessels toflow into and out of the apparatus, as depicted in FIG. 14. Immediateperfusion of oxygenated blood occurs, which allows survival and functionof the entire living mass.

In a one embodiment, tissue-engineered liver is formed. Preferably,tissue engineered liver comprises both functioning hepatocytes and bileducts. (See FIG. 9B.) The biliary system of native liver begins with aminute hexagonal bile canaliculus, which is formed from specializationof the adjacent surfaces of individual hepatocytes, which are sealedwith tight junctions. These canaliculi are confluent with terminalbiliary ductules, which are initially made of squamous cells, but giveway to low cubiodal biliary epithelium as they approach the interlobularbile ducts. One liter of bile per day is secreted by hepatocytes andmoved out of the liver through this system. There have been previousreports of the formation of duct-like structures in a variety oflong-term in vitro and in vivo hepatocyte cultures (Block, et al., JCell Biol, 132, 1133 (1996); Landry, et al., J Cell Biol, 101, 914(1985); Mitaka, et al., Hepatology 29, 111 (1999); Nishikawa, et al.,Exp Cell Res, 223, 357 (1996); Uyama, et al., Transplantation 55, 932(1993)). In Example 4, engineered liver tissue, cultured inmicrofabricated channels of a single layer mold, is shown to comprisestructures resembling bile ducts.

In a yet another embodiment, tissue-engineered kidney is formed.Preferably, tissue engineered kidney comprises functioning proximaltubules. Tissue-engineered kidney functions as a native kidney;glomerular ultrafiltrate flows from the glomerular endothelium andpasses through a semipermeable membrane into a proximal tubule networkwhere reabsorption occurs (see Example 5).

Extracomoreal Support Devices

The invention can be adapted to comprise devices for uses in addition tothe formation of implantable tissue. Such devices can be extracorporeal,and may provide partial support function, may extend the time betweenhospital treatments for patients on chronic organ support therapies, andwill improve the quality of life between hospital treatments. Currentextracorporeal devices known in the art do not incorporate the precisemicrofabrication capabilities offered by MEMS technology, and thereforefunction is limited by the resolution limits of current hollow fiber andmembrane fabrication methods. Insertion of MEMS technology into thisdomain is expected to have major benefits for hemofiltering, dialysisand other applications. For example, the designs can be adapted toproduce an extracorporeal renal dialysis device, an extracorporeal liverdevice, or an extracorporeal artificial lung device. Such devices may ormay not be supported with living cells loaded or seeded into the device.

The systems of the invention can be implanted into a subject tosupplement or replace the biological function of a tissue or organ.Alternatively, the systems can remain ex viva, serving as extracorporealdevices to supplement or replace biological function. As used herein,the term “biological function” refers to the structural, mechanical ormetabolic activity of a tissue or organ. Extracorporeal devices of thepresent invention can comprise hybrid devices suitable for both ex vivoand in vivo use.

The Examples presented herein demonstrate that microfabricationtechnology can be adapted to suit the needs of all living tissues. Animportant aspect of the present invention lies in its control of formover extremely small distances. The resolution of the microfabricationtechniques is on the order of about 0.1 μm from point to point. Thislevel of precision adds new levels of control in the ability to designand guide new tissue formation. For instance, surfaces can be imprintedwith submicron grooves or scallops, and corners can be made rounded,angled or sharp with this same level of submicron precision. Geometriccontrol at this scale can have a powerful impact on cell adhesionthrough mechanisms such as contact guidance, as described by Den Braber,et al. J. Biomed. Mater. Res. 40, 291 (1998).

The following Examples are provided to illustrate the invention, andshould not be considered to limit its scope in any way.

Example 1 Micromaching of Template to Tissue Engineer BranchedVascularized Channels for Liver Fabrication

Micromachining technologies were used on silicon and pyrex surfaces togenerate complete vascular systems that can be integrated withengineered tissue before implantation. Trench patterns reminiscent ofbranched architecture of vascular and capillary networks were etchedusing standard photolithographic techniques onto silicon and pyrexsurfaces to serve as templates. Hepatocytes and endothelial cells werecultured and subsequently lifted as single-cell monolayers from thesetwo dimensional molds. Both cell types were viable and proliferative onthese surfaces. In addition, hepatocytes maintained albumin production.The lifted monolayers were then folded into compact three-dimensionaltissues. The goal is to lift these branched vascular networks fromtwo-dimensional templates so that they can be combined with layers ofparenchymal tissue, such as hepatocytes, to form three-dimensionalconformations of living vascularized tissue for implantation.

Materials and Methods

Micromachining Techniques

Templates for the formation of sheets of living vascularized tissue werefabricated utilizing micromachining technology. For the present work, asingle level etch was utilized to transfer a vascular network patterninto an array of connected trenches in the surface of both silicon andpyrex wafers.

In this prototype, a simple geometry was selected for patterning thevascular network. Near the edge of each wafer, a single inlet or outletwas positioned, with a width of 500 μm. After a short length, the inletand outlet branched into three smaller channels of width 250 μm; each ofthese branched again into three 125 μchannels, and finally down to three50 μn channels. Channels extend from the 50 μm channels to form acapillary network, which comprises the bulk of the layout. In betweenthese inlet and outlet networks lies a tiled pattern of diamonds andhexagons forming a capillary bed and filling the entire space betweenthe inlet and outlet. In one configuration, the capillary width was setat 25 μm, while in the other, capillaries were fixed at 10 μm. Thisgeometry was selected because of its simplicity as well as its roughapproximation to the size scales of the branching architecture of theliver. Layout of this network was accomplished using CADENCE software(Cadence, Chelmsford, Mass.) on a Silicon Graphics workstation. A filewith the layout was generated and sent electronically to Align-Rite(Burbank, Calif.), where glass plates with electron-beam-generatedpatterns replicating the layout geometry were produced and returned forlithographic processing.

Starting materials for tissue engineering template fabrication werestandard semiconductor grade silicon wafers (Virginia Semiconductor,Powhatan, Va.), and standard pyrex wafers (Bullen Ultrasonics, Eaton,Ohio) suitable for MEMS processing. Silicon wafers were 100 mm diameterand 525 μm thick, with primary and secondary flats cut into the wafersto signal crystal orientation. Crystal orientation was <100>, and waferswere doped with boron to a resistivity of approximately 5 W-cm. Thefront surface was polished to an optical finish and the back surfaceground to a matte finish. Pyrex wafers were of composition identical toCorning 7740 (Corning Glass Works, Corning N.Y.), and were also 100 mmin diameter, but had a thickness of 775 μm. Both front and back surfaceswere polished to an optical finish. Prior to micromachining, both wafertypes were cleaned in a mixture of 1 part H₂SO₄ to 1 part H₂O₂ for 20minutes at 140° C., rinsed 8 times in deionized water with a resistivityof 18 MW, and dried in a stream of hot N₂ gas.

For silicon and pyrex wafers, standard photolithography was employed asthe etch mask for trench formation. Etching of pyrex wafers requiresdeposition of an intermediate layer for pattern transfer which isimpervious to the etch chemistry. A layer of polysilicon of thickness0.65 μm over the pyrex was utilized for this purpose. This layer wasdeposited using Low Pressure Chemical Vapor Deposition (LPCVD) at 570°C. and 500 mTorr via the standard silane decomposition method. In thecase of silicon, photoresist alone could withstand limited exposure totwo of the three etch chemistries employed. For the third chemistry, a1.0 μm layer of silicon dioxide was thermally deposited at 1100° C. inhydrogen and oxygen.

Once the wafers were cleaned and prepared for processing, images of theprototype branching architecture were translated onto the wafer surfacesusing standard MEMS lithographic techniques. A single layer ofphotoresist (Shipley 1822, MicroChem Corp., Newton, Mass.) was spun ontothe wafer surfaces at 4000 rpm, providing a film thickness ofapproximately 2.4 μm. After baking at 90° C. for 30 minutes, the layerof photoresist was exposed to uv light using a Karl Russ MA6 (SussAmerica, Waterbury, Vt.) mask aligner. Light was passed through thelithographic plate described earlier, which was in physical contact withthe coated wafer. This method replicates the pattern on the plate to anaccuracy of 0.1 μm. Following exposure, wafers were developed in Shipley319 Developer (MicroChem Corp., Newton, Mass.), and rinsed and dried indeionized water. Finally, wafers were baked at 110° C. for 30 minutes toharden the resist, and exposed to an oxygen plasma with 80 Watts ofpower for 42 seconds to remove traces of resist from open areas.

Silicon wafers were etched using three different chemistries, whilepyrex wafers were processed using only one technique. For pyrex, thelithographic pattern applied to the polysilicon intermediate layer wastransferred using a brief (approximately 1 minute) exposure to SF₆ in areactive-ion-etching plasma system (Surface Technology Systems, Newport,United Kingdom). Photoresist was removed, and the pattern imprinted intothe polysilicon layer was transferred into trenches in the silicon usinga mixture of 2 parts HNO₃ to 1 part HF at room temperature. With an etchrate of 1.7 μm per minute, 20 μm deep trenches were etched into thepyrex wafers in approximately 12 minutes. Since the chemistry isisotropic, as the trenches are etched they become wider. Processing withthe layout pattern with 25 μm wide capillary trenches tended to resultin merging of the channels, while the use of 10 μm wide trenches avoidedthis phenomenon. Interferometric analysis of the channels after etchingshowed that surface roughness was less than 0.25 μm. Once channeletching of pyrex wafers was completed, polysilicon was removed with amixture of 10 parts HNO₃ to 1 part HF at room temperature, and waferswere recleaned in 1 part H₃SO₄ to 1 part HF.

Three different chemistries were employed to etch silicon in order toinvestigate the interaction between channel geometry and cell behavior.First, a standard anisotropic plasma etch chemistry, using a mixture ofSF₆ and C4F₈ in a switched process plasma system from STS²⁴, was used toproduce rectangular trenches in silicon. Narrower trenches are shallowerthan deep trenches due to a phenomenon known as RIE lag. A secondprocess utilized a different plasma system from STS, which producesisotropic trenches with a U-shaped profile. While the process isisotropic, widening of the trenches is not as severe as is experiencedin the isotropic pyrex etching process described earlier. In both ofthese plasma etching cases, trenches were etched to a nominal depth of20 μm. For the third process, anisotropic etching in KOH (45% w/w in H₂Oat 88° C.), the intermediate silicon dioxide layer mentioned above wasemployed. First, the silicon dioxide layer was patterned using HFetching at room temperature. The KOH process produces angled sidewallsrather than the rectangular profile or U-shaped profile produced by thefirst two recipes, respectively. Crystal planes in the <111> orientationare revealed along the angled sidewalls, due to anisotropic propertiesof the KOH etch process as a function of crystal orientation. Due to theself-limiting nature of the channels produced by this process, trenchdepth was limited to 10 μm. After completion of the silicon waferetching, all layers of photoresist and silicon dioxide were removed, andwafers were cleaned in 1 part H₂SO₄:1 part H₂O₂ at 140° C., followed byrinsing in deionized water and drying in nitrogen gas.

For this set of experiments, no attempt was made to alter the surfacechemistry of the silicon and pyrex wafers. Prior to processing, siliconwafers were uniformly hydrophobic, while pyrex wafers were equallyhydrophilic, as determined by observations of liquid sheeting andsessile drop formation. After processing, unetched surfaces appeared toretain these characteristics, but the surface chemistry within thechannels was not determined.

Animals

Adult male Lewis rats (Charles River Laboratories, Wilmington, Mass.),weighing 150-200 g, were used as cell donors. Animals were housed in theAnimal Facility of Massachusetts General Hospital in accordance withguide lines for the care of laboratory animals. They were allowed ratchow and water ad libitum and maintained in 12-hour light and darkcycle.

Cell Isolations

Male Lewis rats were used as hepatic cell donors. HCs were isolatedusing a modification of the two-step collagenase perfusion procedure aspreviously described by Aiken, et al., J Pediatr Surg 25, 140 (1990) andSeglen, Methods Cell Biol 13, 29 (1976). Briefly, the animals wereanesthetized with Nembutal Sodium Solution (Abbott Laboratories, NorthChicago, Ill.), 50 mg/kg, and the abdomen was prepared in sterilefashion. A midline abdominal incision was made and the infrahepaticinferior vena cava was cannulated with a 16-gauge angiocatheter (BectonDickinson), The portal vein was incised to allow retrograde efflux andthe suprahepatic inferior vena cava was ligated. The perfusion wasperformed at a flow rate of 29 ml/min initially with a calcium-freebuffer solution for 5 to 6 minutes, then with a buffer containingcollagenase type 2 (Worthington Biomedical Corp., Freehold, N.J.) at 37°C. The liver was excised after adequate digestion of the extracellularmatrix and mechanically agitated in William's E medium (Sigma, St.Louis, Mo.) with supplements to produce a single cell suspension. Thesuspension was filtered through a 300 μm mesh and separated into twofractions by centrifugation at 50 g for 2 minutes at 4° C. The pelletcontaining the viable HC fraction was resuspended in William's E mediumand further purified by an isodensity Pereoll centrifugation. Theresulting pellet was then resuspended in Hepatocyte Growth Medium, andcell counts and viabilities of HCs were determined using the trypan blueexclusion test.

The endothelial cells were derived from rat lung microvessels and theywere purchased directly from the vendor, Vascular Endothelial CellTechnologies (Rensellaer, N.Y.).

Hepatocyte Culture Medium

William's E medium supplemented with 1 g sodium pyruvate (Sigma, St.Louis, Mo.) and 1% glutamine-penicillin-streptomycin (Gibco BRL,Gaithersburg, Md.) were used during the cell isolation process. Theplating medium was Dulbecco's modified eagle medium (Gibco BRL)supplemented with 10% fetal bovine serum, 1% penicillin-streptomycin, 44mM sodium-bicarbonate, 20 mM HEPES, 10 mM niacinamide, 30 microgram/nilL-proline, 1 mM ascorbic acid 2 phospate, 0.1 μM dexamethasone (Sigma),insulin-transferrin-sodium selenite (5 mg/L-5 mg/L-5 μgram/L, RocheMolecular Biomedicals, Indianapolis, Ind.), and 20 rig/mL epidermalgrowth factor (Collaborative Biomedical Products, Bedford, Mass.).

Endothelial Cell Culture Medium

Dulbecco's modified eagle medium (Gibco BRL) was supplemented with 10%fetal bovine serum, 1% penicillin-streptomycin, 25 mg of ascorbic acid(Sigma), 10 mg L-alanine (Sigma), 25 mg L-proline (Sigma), 1.5 microgramcupric sulfate (Sigma), glycine (Sigma) and 1M Hepes buffer solution(Gibco BRL). The media was supplemented with 8 mg of ascorbic acid everyday.

Cell Attachment and Lifting from Non-Etched Silicon and Pyrex Wafers

Silicon and pyrex were both tested as possible substrates for theculture and lifting of endothelial cells and hepatocytes. Prior to cellseeding, the pyrex wafers were sterilized with 70% ethanol (Fisher,Pittsburgh, Pa.) overnight and washed three times with sterile phosphatebuffered saline (Gibco BRL). Silicon wafers were first soaked in acetonefor 1 hr, followed a methanol rinse for 15 minutes, and overnightsterilization in 100% isopropyl alcohol. Rat lung microvascularendothelial cells was cultured on non-coated pyrex and silicon surfaces,as well as wafers coated with vitrogen (30 microgram/ml), Matrigel®(1%), or Gelatin (10 mg/ml). Once isolated, the cells were resuspendedin endothelial cell culture medium, seeded uniformly onto the wafer at adensity of 26.7×10³ cells/cm², and cultured at 5% CO₂ and 37° C. Afterreaching confluence, the ability of the monolayer of endothelial cellsto lift from the wafers was tested using a cell scrapper to promotedetachment.

The rat hepatocytes were also cultured on non-coated pyrex and silicon,as well as wafers coated with a thin and thick layers of vitrogen (30microgram/ml and 3 microgram/ml) and Matrigel (1%) in order to determinethe optimal methods for lifting hepatocyte sheets. Once isolated, thehepatocytes were resuspended in hepatocyte growth media, seeded onto thewafer at a density of 111.3×10³ cells/cm², and cultured at 5% CO₂ and37° C. Cell attachment and growth was observed daily using microscopyand cell lifting occurred spontaneously.

After determining which method for culturing was best for lifting thehepatocytes and endothelial cells in an intact layer, both membraneswere fixed in 10% buffered formalin for 1 hr and harvested forhistological study, and the hepatocytes were stainedimmunohistochemically.

Immunohistochemical Staining

The hepatocyte cell monolayer membrane was fixed in 10% bufferedformalin and processed for hematoxylin-eosin and immunohistochemicalstaining using a labeled streptavidin biotin method (LSAB2 kit for ratspecimen, DAKO, Carpinteria, Calif.). The primary antibody was rabbitanti-albumin (ICN, Costa Mesa, Calif.). Three-micron sections wereprepared and deparafinized. The specimens were treated with peroxidaseblocking buffer (DAKO) to prevent the nonspecific staining. Sectionswere stained with albumin diluted with phosphate buffered saline,followed by biotinylated anti-rabbit antibody and HRP conjugatedstreptavidin. Sections were treated with DAB as substrate and werecounterstained with hematoxylin.

Albumin Production

To assess hepatocyte function, albumin concentration in the culturemedium was measured every 24 hours for 5 days pre-cell detachment usingan enzyme linked immunosorbent assay (n=5), as described by Schwere, etal., Clinica Chernica Acta 163, 237 (1987). In brief, a 96 wellmicroplate was coated with anti-rat albumin antibody (ICN). Afterblocking non-specific responses with a 1% gelatin solution, each samplewas seeded onto the plate and incubated for 1 hour at 37° C. This wasfollowed by another 1 hour incubation with peroxidase conjugatedanti-rat albumin antibody (ICN). Finally, the substrate was added andextinction was measured with a microplate reader at 410 nm. R² of thestandard curve was >0.99. Results demonstrate continued production ofalbumin by cultured hepatocytes (FIG. 15).

Statistical Analysis

All data was expressed as mean+/−SD. Statistical analysis was performedwith a paired t-test. Statistical significance was determined as whenthe p value of each test was less than 0.05.

Cell Attachment to Etched Silicon and Pyrex Wafers

Endothelial cells and hepatocytes were also seeded onto etched siliconand pyrex wafers. Prior to cell seeding, the pyrex wafers weresterilized with 70% ethanol (Fisher) overnight and washed three timeswith sterile phosphate buffered saline (Gibco BRL). Silicon wafers werefirst soaked in acetone for 1 hr, followed a methanol rinse for 15minutes, and overnight sterilization in 100% isopropyl alcohol. Ontothese wafers were seeded rat lung microvascular endothelial cells at adensity of 26.7×10³ cells/cm², or rat hepatocytes at a density of111.3×10³ cells/cm². These cells were cultured at 5% CO² and 37° C., andtheir attachment and growth observed daily using microscopy.

Implantation of Hepatocyte Sheets into the Rat Omentum

Hepatocytes were cultured on silicon wafers coated with a thin layer ofvitrogen (30 microgram/rill), and lifted in sheets. Retrorsine is a drugknown to inhibit the regeneration of the normal liver by producing ablock in the hepatocyte cell cycle with an accumulation of cells in lateS and/or G₂ phase (Peterson J E J Pathol Bacterial 89, 153 (1965)). Thisdrug was administered into the peritoneal cavity of two rats at a doseof 3 mg/ml/100 g on day 0, and after two weeks. Three weeks later, aportacaval shunt was created, and the following week a hepatocyte sheet,lifted after four days culture on vitrogen coated silicon (30microgram/ml), was implanted onto the microvasculature of the ratomentum and rolled into a three-dimensional cylinder, and a 60%hepatectomy was performed. The rolled omentum with hepatocytes washarvested at four weeks and at three months after implantation andanalyzed using histology.

Results

Micromachining

A schematic of the vascular branching network design used as a templatefor micromachining is shown in FIG. 16A. This pattern was transferred tosilicon and pyrex wafers using the processes described in the Materialsand Methods section. Typical trench depths of 20 μm on silicon and 10 μmon glass were achieved utilizing these processes. An optical micrographof a portion of the capillary network etched into a silicon wafer isshown in FIG. 16B. In FIG. 16C, a Scanning Electron Micrographcross-section of an angled trench etched using the anisotropic etchingprocess described earlier is shown. This process resulted in excellentadhesion and enhanced lifting of living tissue.

Growth and Lifting of Cells from the Silicon and Pyrex Wafers

The adhesion and growth of endothelial cells and hepatocytes on severaldifferent substrate surfaces were compared. On all pyrex wafers, coatedor non-coated, the endothelial cells proliferated and grew to confluencewithin four days. These cells did not lift spontaneously, and whenscraped, did not lift as a single sheet. In addition, when thenon-coated silicon wafers were seeded with endothelial cells, the cellsheet fragmented upon lifting. On the other hand, endothelial cellsseeded onto silicon surfaces coated with vitrogen (30 microgram/ml),Matrigel (1%), and gelatin (10 mg/ml) did lift with the use ofmechanical means (i.e. a cell scraper), and provided an intact monolayersheet of endothelial cells. Upon observation, there were no significantdifferences in the effects of the three coatings on the detached cellsheets.

Hepatocytes also attached and spread well on all coated and non-coatedpyrex wafers, and did not lift spontaneously or in sheets when scrapedafter several days of growth. However, when seeded onto silicon wafers,they lifted spontaneously on all the non-coated and coated wafers. Thehepatocyte sheets lifted from the non-coated wafers after 3 days, butwere very fragile and fragmented easily. The monolayers that lifted fromthe thin and thickly coated vitrogen substrates (30 microgram/ml and 3microgram/ml) lifted after 4 days in culture to form an intacthepatocyte layer. Cells lifted from the Matrigel (1%) coated siliconwafers after 5 days in culture. There were no significant differences inappearance between the cell sheets lifted from the vitrogen and Matrigelcoated wafers.

Histological assessment of the detached cell monolayers of bothhepatocytes and endothelial cells manifested promising results.Hernotoxylin and Eosin (H&E) staining of both showed that all cells wereviable and that most were undergoing mitoses. The endothelial cells wereobserved to be primarily attenuated and to form a single-celledalignment. The monolayer of hepatocytes showed each cell to be of aspheriod configuration with eosinophilic floculent cytoplasm and a largenucleus with a bright red nucleolus, similar to that seen in the nativeliver. Moreover, cellular attachments were less attenuated than theendothelial cells. Thus, these results are reminiscent of each of thecell types' specific functions. In biological systems, the endotheliumfunctions to provide a thin, smooth outer surface of a barrier and atransport channel and so it is understandable that these cells areobserved here to be primarily attenuated and in a single-celled array.The hepatocytes have more of a tendency to form tissue and so less of asingle-celled array and more of a rounded multi-layered array is seen.

Albumin secretion into the hepatocyte culture medium at day 2, 3, 4, and5 was 165.96±29.87, 164.44±17.22, 154.33±18.46, 115.47±18.09(microgramiday, Graph 1), respectively. Though there was a statisticallysignificant difference between day 4 and day 5, no significantdifferences were observed between day 2, day 3, and day 4 (p<0.05 by thepaired t-test). Hence, this data shows that cells cultured on siliconwafers were able to maintain a fairly constant albumin production rateuntil day 4.

Moreover, through immunohistochemical staining of the detachedhepatocyte monolayers, many cells were stained positive for albuminindicating further that hepatocyte function was maintained on siliconwafers.

Implantation of Hepatocyte Sheet into the Rat Omentum

H&E staining of hepatocyte sheets implanted into rat omentumdemonstrated that all cells were viable and showed proliferation at fourweeks and three months. The implanted hepatocyte monolayer sheets, whenharvested, were over 5 cell layers thick in most areas. This studydemonstrates that silicon microfabrication technology can be utilized toform large sheets of living tissue. It also demonstrates the feasibilityof etching ordered branching arrays of channels that allow livingendothelial cells to line the luminal surface of the channels. Inaddition, it has been shown that organized sheets of engineeredhepatocyte tissue and endothelial tissue can be lifted from the surfaceof silicon or pyrex wafers and can be folded into a compactthree-dimensional configuration. The hepatocyte sheets have then beenplaced into rats on the highly vascular surface of the omentum. Thatstructure has then been rolled into a three-dimensional cylinder as amodel for an engineered vasculature. Vascularized hepatic tissue wasformed as a permanent graft.

Example 2 Endothelialized Microvascular Networks Grown on MicromachinedPyrex® Templates for Tissue Engineering of Vital Organs

This Example shows the design, modeling, and experimental/computationaltesting of the endothelialized microvascular matrices grown onmicromachined Pyrex® templates.

Patterns of microvascular networks were etched using microfabricationtechnologies on Pyrex® wafers. The pattern consisted of 10 generationsof bifurcations from a single inflow channel of width 3 mm down tochannels of width of 30 μm.

The channels were then collected to a single outflow. All channels wereetched to the same depth of 30 μm. The Pyrex® wafer was sealed to a flatsilicone rubber sheet of the same size. Endothelial cells harvested fromrat lung were successfully seeded and expanded under continuous flowconditions in this microvascular network. Red blood cells harvested fromrat were heparinized and perfused into the endothelialized channels, andsuccessfully collected at the output. Using micro-visualizationtechniques, the concentration of red blood cells (hematocrit) in themicrovascular network was measured. The distribution of blood flow rate,pressure, and hematocrit was also calculated in the entire microvascularsystem using an earlier developed computational algorithm.

Epithelial cells were observed flowing through channels and attachingmainly around the walls of smallest channels on day 1 and growing toconfluence along the channels under continuous flow conditions over thefollowing 5 days. Rat lung endothelial cells attach in a single layer tothe walls of these mold structures without occluding them.

Hematocrit compared well between the experimental measurements andnumerical calculations. Red blood cells reach even the smallest vesselsin the network, ensuring sustained transport of oxygen to the engineeredcapillaries.

In summary, microfabrication technology is demonstrated as an approachfor organizing endothelial cells in vitro at the size scale of themicrocirculation.

Example 3 Microfluidics Device for Tissue Engineering Microvasculature:Endothelial Cell Culture

In this Example, the fabrication of the microfluidic mold, in vitroseeding, and extended cell culture in the mold is demonstrated.Capillary networks were fabricated in biocompatible PDMS, sterilized,coated with cell adhesion molecules, and seeded with cells.Cell-containing molds were then connected to a closed-loop bioreactorfor long-term culture. Continuous-flow culture of endothelial cells forup to 4 weeks without occlusion or contamination was achieved.

Traditional soft lithography microfluidics were used as a prototypematrix. These cell-containing microfluidics are capable of supportinglong-term culture in vitro, because in vitro expansion of cells prior toimplantation can take several weeks. The prototype matrix is designed tosupply sufficient oxygen and nutrients and to remove excretory productswhile avoiding large shear stresses. The matrix is useful for long-termmicrofluidic cell culture, including the maintenance of sterility andthe minimization of cell and bubble occlusions.

Microfluidic networks that support physiologic flows and pressures weredeveloped by photopatterning SU-8, a high-aspect ratio negativephotoresist, onto silicon. This was used as a mold for castingpolydimethylsiloxane (PDMS). After removal from the mold, inlets andoutlets were cored with blunted syringe needles, and the micropatternedpolymer scaffold was irreversibly sealed to an unpatterned layer ofPyrex® or PDMS by oxygen plasma surface treatment. See Duffy, et al.,Anal. Chem. 70, 4974 (1998). The microfluidic device was autoclavesterilized and perfused with a solution containing cell adhesionmolecules (poly-L-lysine, collagen, gelatin, or fibronectin), which wereallowed to adsorb for one hour.

The fluidic network was then seeded with a 1×10⁶-1×10⁸ cells/mL cellsuspension using a syringe pump at flow rates ranging from 10-100μL/min. The cells were then allowed to attach for 24 hours, after whichthe device was connected in-line with a sterile bioreactor consisting ofa peristaltic pump, oxygenator, bubble trap, and a reservoir of sterileculture medium. Sterile culture medium was pumped peristaltically from asterile reservoir through an oxygenator consisting of along length oftubing semipermeable to oxygen. The oxygenator was followed by a smallbubble trap, leading directly to the microfluidic circuit. Finally, thesystem was run closed-loop in an incubator at standard culture settings.

Autoclave sterilization of the microfluidic circuit caused no obviouspattern distortion. Coating the channels with cell adhesion moleculesenhanced cell attachment when compared to phosphate bufferedsaline-coated control channels. Seeding of cells into channels of widthsbetween 30-200 μm was optimized by varying concentrations and flowrates. The continuous-flow bioreactor was used to dynamically cultureendothelial cells at flow rates between 0.01 mL/min and 0.1 mL/min. Bothsingle channels and complex networks of channels (30-200 μm wide and 40μm deep) were successfully seeded and cultured. In 100 μm×40 μm singlechannels, cells were cultured for more than 4 weeks withoutcontamination or occlusion.

Long term culture of cells in microfluidic devices was achieved. Cellssuccessfully attached, proliferated, and migrated in closedmicrofabricated channels with small geometries.

Example 4

Generation of Functionally Differentiated, Three-Dimensional HepaticTissue from Two-Dimensional Sheets of Small Hepatocytes andNon-Parenchymal Cells

In this Example, three-dimensional, vascularized liver tissue wasfabricated in vivo from a non-vascularized monolayer or cell sheet ofsmall hepatocytes (SHCs) formed on a silicon wafer. SHCs cells aresmaller than mature hepatocytes (MHCs), but morphologically similar,with a highly proliferative capacity (Mitaka, et al., Biochem BiophysRes Commun 214, 310 (1995); Mitaka, et al., Gastroenterol Ilepatol 13Suppl, S70 (1998); Mitaka, et al., Hepatologoy, 29, 111 (1999); Tateno,et al., Am J Pathol 148, 383 (1996); Tateno, et al., Am J Pathol 149,1593 (1996)).

Cell sheets created from SHCs and NPCs were implanted onto rat omentumwith maximal hepatotrophic stimulation by retrorsine, portacaval shunt,and partial hepatectomy, and their engraftment and function wereevaluated. Using this cell type, co-cultured with nonparenchymal cells(NPCs), liver tissue that maintained a high level of albumin productionwas fabricated in a flow culture system. Animals as described in Example1 were used as cell donors. Cells were cultured as described in theHepatocyte Culture Medium section of Example 1.

Cell Isolation

SHCs and NPCs were isolated by the process described in Example 1, withthe following modifications. Animals were anesthetized by anintramuscular injection with Ketamine and Xylazine. Cells werecollected, suspended, filtered and centrifuged as previously described.Following centrifugation, the pellet containing a majority of MHCs wasdiscarded. The supernatant was collected, and the fraction containingSHCs and NPCs was obtained as a pellet by additional centrifugationtwice at 150×g for 5 minutes. The pellet was resuspended in the platingmedium and the cell number and viability were counted using the trypanblue exclusion test.

In Vitro SHC Sheets Preparation

In order to obtain SHC sheets, the SHCs and NPCs were seeded andcultured on silicon wafers (10 cm diameter). Briefly, the silicon waferswere sterilized with ethylene oxide gas and coated with liquid collagen(Vitrogen 100, Collagen Corp., Palo Alto, Calif.). The mixture of SHCsand NPCs was resuspended in the plating medium at a density of 0.8×10⁶cells/mL. A 25 mL suspension was seeded onto the silicon wafer in a 15cm Petri dish and incubated at 37° C., 5% CO₂. The plating medium waschanged every other day. After reaching confluence, the cultured cellswere lifted as sheet by scraping with a sterile razor blade and preparedfor implantation.

Albumin Production

To assess SEC function before implantation, albumin concentration in theplating medium was measured at 3, 5, 7 and 10 days after cell seedingusing an enzyme linked immunosorbent assay (ELISA) (n=11) as describedin Example 1.

In Vivo Model

Retrorsine was administered into the peritoneal cavity of recipient rats(n=23) at a dose of 3 mg/ml/100 g on day 0, and after two weeks aspreviously reported (Laconi, et al., Am J. Pathol 153, 319 (1998)).Three weeks after the second administration, an end-to-side portacavalshunt was created using 8-0 Ethilon suture (ETHICON, Somerville, N.J.)to generate systemic hepatoirophic stimulation for SHC sheetimplantation. One week later, a SHC sheet was spread onto the ratomentum and rolled from distal to proximal into a three-dimensionalcylinder. The omentum was sutured to the anterior wall of the stomachusing 7-0 Prolene suture (ETHICON). A 60% partial hepatectomy wasperformed simultaneously for hepatotrophic stimulation. Animals weresacrificed at the designated time points after SHC sheet implantationfor specimen retrieval. The resected specimens were fixed in 10%formalin solution (Sigma), routinely processed and embedded in paraffinfor subsequent hematoxylin-eosin (H & E) and immunohistochemicalstaining. Two specimens were fixed in 2.5% gluteraldehyde (Sigma) forelectron microscopy (EM).

Immunohistochemical Staining

To characterize the implanted constructs, immunohistochemical stainingusing the Avidin-biotin peroxidase complex (ABC) method was performed.The primary antibodies included: rabbit anti-albumin (DAKO, Carpinteria,Calif.), rabbit anti-transferrin (ICN), mouse anti-pancytokeratin(Sigma), goat anti-γ-glutamyl transpeptidase (OTT) (a gift from Dr.Petersen, Department of Pathology, University of Florida, Fla.). Four.mu.m paraffin sections were deparaffinized and treated with 4.5% H₂O₂in methanol. The specimens were digested for 12 minutes with 0.1%trypsin solution, followed by treatment with avidin D (Vector) and 5%serum. Subsequently, slides were incubated with the respective primaryantibody that were diluted in phosphate buffered saline with 1% bovineserum albumin overnight at 4° C. Biotinylated antimouse/rabbit/goatantibody was used as a secondary antibody in combination with theVectastain ABC kit (Vector, Burlingame, Calif.). Finally, specimens weretreated with 3-amino-9-ethylcarbazole (AEC) (Vector) as substrate andwere counterstained with. Mayer's hematoxylin solution (Sigma).

Electron Microscopy (EM)

Two rats at 4 months were sacrificed for EM study. Immediately afterremoval from the animal, 1 mm sections were placed into Kamovsky's KIIsolution (2.5% glutaraldehyde, 2.0% paraformaldehyde, 0.025% calciumchloride, in a 0.1 M sodium cacodylate buffer, pH 7.4), fixed overnightat 4° C., and routinely processed for EM. Representative areas werechosen from 1 μm sections stained with toluidine blue. The sections wereexamined using a Phillips 301 transmission electron microscope.

Morphologic and Quantitative Analysis

For morphologic and quantitative analysis, specimens were harvested at 2weeks (n=7), 1 month (n=7), and 2 months (n=7). At each time point, therolled omentum was cut perpendicularly to the greater curvature ofstomach, and three to four cross-sections of tissue were obtained andstained with H & E. The area occupied by implanted constructs in eachsection was measured using computer assisted analysis with NIH Imageversion 1.61 software (Division of Computer Research and Technology,NIH, Bethesda, Md., USA). This was expressed as μm²/section.

Statistical Analysis

All values are expressed as mean f SD and were statistically evaluatedusing the Mann-Whitney test or the paired t-test. A value of p<0.05 wasconsidered statistically significant.

Cell Isolation and Growth in a Culture Flask

All cell isolations yielded 8-14×10⁷ cells comprising SHCs and NPCswith >90% overall viability. To evaluate the culture condition of SHCson silicon wafers, a cell suspension was seeded on culture flasks at thesame concentration. One day after seeding, most cells began to attachindividually or occasionally form small clusters consisting of severalSHCs. After 3 days, cells have completely attached and spread on theculture flask. After 5 days, many clusters had formed and NPCs wereobserved between the clusters. These small clusters united to formlarger clusters and continued to grow until implantation (FIG. 17).

Cell Sheet Formation

SHCs and NPCs cultured on silicon wafers grew similarly to the cultureflask. Many large clusters were observed macroscopically on the siliconwafers after culturing for 10-14 days. Cultured cells were lifted as asheet from all the silicon wafers. After lifting, cell sheets shrunk toapproximately 2.5 cm in diameter (FIG. 18)

Albumin Production

Albumin secretion at day 3, 5, 7, and 10 was, 6.47±2.49, 12.08±5.18,19.93±4.05, 30.14±5.46 (au/day), respectively (FIG. 19). There werestatistically significant differences between day 3 and day 5, day 5 andday 7, and day 7 and day 10 (p<0.05 by the paired t-test).

H & E Staining

The H & E staining of specimens harvested at 2 weeks after cell sheetimplantation typically reveal large, polygonal, eosinophilic cells withround nuclei resembling hepatocytes, cuboidal cells resembling biliaryepithelial cells, and capillary formation. At this time point the areaof hepatocytes was less than five cell layers thick (FIG. 20A). At 1 and2 months, large clusters of hepatocytes over five cell layers thick,cuboidal cells resembling biliary epithelial cells, and capillaryformation could be observed (FIGS. 20B-D). In some areas, hepatocytesexceeded ten cells layer thick. In the specimens at 2 weeks and 1 month,there were many areas that were occupied mainly by bile duct-likestructures (FIG. 20D). As the implant matured in the omentum, the numberof hepatocytes increased and the number of bile duct-like structuresdecreased at 2 months.

Immunohistochemistry

Both hepatocytes and bile duct-like structures stained positively withpan-cytokeratin. However, bile ductules stained more strongly positivethan hepatocytes (FIG. 21A). Since there are normally no pan-cytokeratinpositive cells in the omentum, it is likely that the cells originatedfrom the implanted constructs. Some of the hepatocytes stainedpositively for albumin and transferrin, which suggests that theycontinued to express liver specific functions. The bile duct-likestructures stained positively for GGT, an enzyme expressed at highlevels in normal rat intrahepatic biliary epithelial cells but typicallynot detected in normal rat hepatoeytes, and negatively for albumin andtransferrin, which indicated that they were composed of cells resemblingnormal biliary epithelial cells (FIGS. 21B-D).

In one case, histology showed that one bile duct-like structure at 2weeks was formed with both cells resembling biliary epithelial and cellswhich were morphologically more similar to hepatocytes (FIG. 22). Thisbile duct-like structure was located between the canaliculi-likestructures composed of hepatocytes, and the bile duct-like structuresformed solely by cells resembling biliary epithelial as if it were atransitional structure between the two. This phenomenon demonstratesthat canaliculi-like structures and bile duet-like structures grow toconfluence in tissue engineered constructs.

Ultrastructure of the Implanted Construct

TEM revealed that the engineered constructs were composed of cells withlarge round nuclei, numerous mitochondria and peroxisomes, andmicrovilli; characteristic of hepatocytes. These cells formed structuresresembling bile canaliculi at the cell-cell borders. Capillaries wereseen between hepatocytes (FIG. 23B).

Morphologic and Quantitative Analysis

The calculated areas occupying implanted constructs were 43136+/−36181,153810+/−06422, and 224332+/−142143 μm²/section at 2 weeks, 1 month, and2 months, respectively. The mean area increased over time, and therewere significant differences between 2 w and 1 m (p<0.05), and between 2w and 2 m (p<0.01). No significant difference was observed between 1 mand 2 m. The areas occupied by bile duct-like structures were13407+/−16984, 15430+/−8980, and 1290+/−2052 μm²/section at 2 weeks, 1month, and 2 months, respectively. The areas were significantly greaterat 2 weeks (p<0.01) and 1 month (p<0.05 by the Mann-Whitney U test),compared to the area at 2 months (FIG. 24).

This Example shows morphologically simple cell sheets created from SHCsand NPCs implanted and engrafted in the omentum. Given adequatehepatotrophic stimulation, implants formed morphologically complexthree-dimensional tissue consisting of hepatocytes, structuresresembling bile canaliculi, and ducts composed of cells resemblingbiliary epithelium. These results represent a significant advance towardthe tissue engineering of complex vascularized thick tissues.

Example 5 Generation of a Renal Replacement Device

This Example describes a microfabricated network of proximal tubulesthat could potentially replace the essential reabsorptive and excretoryfunctions of the kidney. (See FIGS. 25-28.) A glomerular endothelialcell-lined network can provide filtration while minimizing thrombosis.These two networks combined on bioresorbable polymer are the basis far atissue engineered renal replacement device.

Background

Although the kidney is a complex organ with an intricate vascular supplyand at least 15 different cell types, the critical functions offiltration, reabsorption and excretion can be targeted with tissueengineering. The basic functional unit of the kidney, the nephron, iscomposed of a vascular filter, the glomerulus, and a resorptive unit,the tubule. Filtration is dependent on flow and specialized glomerularendothelial cells. The majority (50-65%) of reabsorption is performed bythe proximal tubule cells using active sodium transport through theenergy-dependent Nat-K⁺-ATPase located on the basolateral membrane. Only5-10% of the approximately one million nephrons in each human kidney isrequired to sustain normal excretory function.

The design of a tissue engineered renal replacement device can then befocused an the development of a glomerular endothelial filter inconjunction with a proximal tubule device for reabsorption andexcretion. The endothelial filter is specifically designed to providephysiologic flow with low thrombogenicity and maximized surface area forsolute transport. The proximal tubule device, containing an appropriatenumber of cells for renal replacement, has optimized surface area forsolute reabsorption and an outlet for urine excretion. (See FIG. 27.)Several layers of molds and/or polymer scaffolds and semi-permeablemembranes can be stacked to optimize filtration and reabsorption (FIG.28). Biocompatible, bioresorbable and microporous polymers are usedthroughout for optimal cell growth and function.

Materials and Methods

Configuring the Mold

MEMS replica molding was used to create the polymer molds used in thisExample (FIG. 2). Using the techniques described herein, an inversepattern (i.e., protrusions rather than indentations) corresponding tothe desired pattern of microchannels was formed on a silicon wafer.Poly-(dimethyl siloxane) (PDMS) was then cast onto the silicon template.After the template was removed, the PDMS was subjected to O₂ plasmatreatment, and was fastened to a second layer of PDMS. In this Example,the second layer of PDMS was flat, however, in other embodiments, eitheror both surfaces of the second PDMS layer can contain a pattern ofmicrochannels. In addition, a semi-permeable membrane can be fastenedbetween the PDMS layers.

Cell Culture

Renal proximal tubule cells and glomerular endothelial cells from ratand pig models have been isolated using sieve filtration and separationover a Percoll gradient (Vinay, et al. Am J Physiol 241, F403 (1981);Misra, et al. Am J Clin Path 58, 135 (1972)). Human microvascularendothelial cells were isolated from normal neonatal foreskin incollaboration with Dr. Michael Detmar (Cutaneous Biology ResearchCenter, MGH Charletown), and stained positively for endothelial cellmarkers CD-31 and von Willebrand's factor (vWF) within the PDMS devices.

Both renal proximal tubule cells and human microvascular endothelialcells were seeded into the MEMS-designed PDMS (poly(dimethyl siloxane))devices at 20 million cells/ml. Cells were allowed to adhere at 37° C.for six hours. Devices were rotated 180° degrees at three hours to allowadherence of cells to both side's of the microchannels. Flow was thenstarted via infusion pump with appropriate culture medium to maintaincell viability.

Animal Model

The appropriate animal is made uremic via bilateral nephrectomies andconnected to a hemoperfusion circuit, The tissue engineered renalreplacement device is connected such that the venous blood enters theglomerular endothelial network and is returned to the animal. The fluidfiltered through the glomerular network then passes through the proximaltubule network. Reabsorbed fluid is returned to the animal, while thefluid remaining in the proximal tubule lumen is analyzed as processedultrafiltrate (urine). The goal rate of hemofiltration is 15-20 ml/min.to match the rate used in renal dialysis. Function of the renalreplacement device is assessed as compared to matched controls forelectrolytes, blood urea nitrogen and creatinine levels, glutathionereabsorption, ammonia excretion, and 1,25-(OH), D₃ levels.

Results

Human microvascular endothelial cells were seeded into poly (dimethylsiloxane) (PDMS) microchannels (smallest channel width 30 depth 35 μm)using the specifically designed MEMS templates, and good cell adherenceand proliferation within the channels was observed (FIG. 29).

A computational model is used to maximize blood flow through theglomerular cell filter, within normal hernodynamic parameters. FiniteElement Modeling (FEM) technologies are used to maximize the surfacearea for filtration to simulate mass transport of solutes across thefilter. The template topography and branching angles are designed tominimize thrombosis within the microchannels. Similarly, the proximaltubule network is optimized to provide even flow distribution, surfacearea for reabsorption, and an outflow tract for excretion of urine.

Cultured proximal tubular cells exhibit characteristic dome formation.Glomerular endothelial cells have also been isolated and maintained inculture. Further characterization of the cells is performed usingimmunohistochemical staining. Proximal tubule cells are stained formegalin (gp330) expression, and endothelial cells are stained for vonWillebrand's factor (vWF) and CD-31.

Function of proximal tubule cells is assessed with the conversion of1,25-OH-D₃ to 1,25-(OH)₂D₃ (1,25-dihydroxyvitamin D₃), the reclamationof glutathione and the generation of ammonium using a single passperfusion system. 25-(OH) D3-12-hydroxylase is a cytochrome P-450monooxygenase found in the inner mitochondrial membrane of proximaltubule cells. Proximal tubule glutathione reclamation is performed bythe brush-border enzyme gamma-glutamyl transpeptidase. In addition,specific transport functions such as vectoral fluid transport (inhibitedby ouabain, an Na⁺-K⁺-ATPase inhibitor), active bicarbonate and glucosetransport (inhibited by acetazolamide and phlorizin respectively), andpara-aminohippurate secretion (inhibited by probenecid) are also tested(Humes, et al., Kid Int 55, 2502 (1999); Humes, et al. Nat Biotechnol17, 451 (1999)). Glomerular endothelial cell function is assessed forpermeability to water and serum proteins, and the basement membranecomponents analyzed.

Microvascular endothelial and proximal tubule cells into have beensuccessfully seeded into PDMS networks made from MEMS templates. FIGS.30-32 show proximal tubule cells growing in the microchannels of thepolymer scaffold at various intervals after seeding.

After generation of microporous biodegradable polymer templates,specific cell attachment and proliferation are tested using DNA or MITassays. Cells on polymer templates are examined for confluence usinghistology and electron microscopy, as well as insulin leak rates (<10%infused). Enhanced attachment of proximal tubule and glomerularendothelial cells to polymers is optimized by precoating the polymersurface with several extracellular matrix components (Matrigel,collagen, fibronectin and laminin) or peptide sequences such as RGD, asdescribed herein.

Flow studies are performed in glomerular endothelial and proximal tubulenetworks in vitro to simulate physiologic blood flow and hemodynamicparameters and to examine cell viability and function.

Having thus described in detail preferred embodiments of the presentinvention, it is to be understood that the invention defined by theappended claims is not to be limited to particular details set forth inthe above description, as many apparent variations thereof are possiblewithout departing from the spirit or scope of the present invention.Modifications and variations of the method and apparatuses describedherein will be obvious to those skilled in the art, and are intended tobe encompassed by the following claims.

1-271. (canceled)
 272. A lamina assembly comprising: a lower structurehaving an upper side defining lower structure channels; an upperstructure having a lower side defining at least one upper structuremicrochannel that opposes the lower structure channels; a semi-permeablemembrane disposed between the lower and upper structures to separate therespective channels; and cells cultured on at least one side of thesemi-permeable membrane.
 273. A lamina assembly as recited in claim 272,wherein the lower and upper structures are fastened together.
 274. Alamina assembly as recited in claim 272, wherein each of the first andsecond structures and the semi-permeable membrane are comprised ofmaterial that is suitable for attachment and culturing of animal cellsand the cells are animal cells.
 275. A lamina assembly as recited inclaim 272, wherein the cells are cultured on both sides of thesemi-permeable membrane.
 276. A lamina assembly as recited in claim 272,wherein the lower structure channels form an inlet in fluidcommunication with an outlet through a plurality of branching channels.277. A lamina assembly as recited in claim 276, further comprising anutrient supply connected to the inlet and an excretion removal lineconnected to the outlet.
 278. A lamina assembly as recited in claim 276,further comprising a pump connected to the inlet for circulating fluidthrough the tissue lamina assembly.
 279. A lamina assembly as recited inclaim 276, wherein the inlet is directly connected to blood vessels ofan animal.
 280. A lamina assembly as recited in claim 272, wherein theupper structure at least one microchannel forms a branching patternidentical to a branching pattern in the lower structure.
 281. (canceled)282. A lamina assembly as recited in claim 272, wherein the upperstructure has an upper side defining upper structure channels, andfurther comprising: a second upper structure having a lower sidedefining second upper structure channels that oppose the upper structurechannels of the upper side; and a second semi-permeable membranedisposed between the upper and second upper structures to separate therespective channels; and cells cultured on at least one side of thesecond semi-permeable membrane.
 283. A lamina assembly as recited inclaim 282, wherein the second upper structure has an upper side definingsecond upper structure channels, and further comprising: a third upperstructure having a lower side defining third upper structure channelsthat oppose the second upper structure channels of the upper side; and athird semi-permeable membrane disposed between the second and thirdupper structures to separate the respective channels; and cells culturedon at least one side of the third semi-permeable membrane.
 284. A laminaassembly as recited in claim 272, wherein the lower structure channelsare open-faced channels and open portions of the lower structurechannels are adjacent to the semi-permeable membrane.
 285. (canceled)286. A lamina assembly as recited in claim 272, further comprising asecond tissue lamina assembly attached to the upper structure, thesecond tissue lamina assembly including: a third structure having a sidedefining third structure channels; a fourth structure having a sidedefining fourth structure channels that oppose the third structurechannels; a second semi-permeable membrane disposed between the thirdand fourth structures to separate the respective channels; and cellscultured on at least one side of the semi-permeable membrane.
 287. Alamina assembly for supplementing or replacing at least one organfunction, comprising: a lower structure having an upper side defining aplurality of microchannels; an upper structure having a lower sidedefining at least one channel that opposes at least a portion of themicrochannels; and a semi-permeable membrane disposed between themicrochannels and the at least one channel.
 288. A lamina assembly asrecited in claim 287, wherein the plurality of microchannels haveendothelial cells seeded therein and the at least one channel hasparenchymal cells therein.
 289. A lamina assembly as recited in claim287, wherein the semi-permeable membrane allows exchange of oxygen,nutrients and waste between fluid circulating in the plurality ofmicrochannels and the at least one channel.
 290. A lamina assembly asrecited in claim 287, wherein each of the first and second structuresand the semi-permeable membrane are comprised of material that issuitable for attachment and culturing of animals cells and the cells areanimal cells and the cells are cultured on both sides of thesemi-permeable membrane.
 291. A lamina assembly as recited in claim 287,wherein the lower and upper structures are fastened together and theplurality of microchannels form at least one inlet in communication withat least one outlet.
 292. A lamina assembly as recited in claim 287,wherein the lower and upper structures are fastened together and theplurality of microchannels form at least one inlet in communication withat least one outlet.
 293. A lamina assembly as recited in claim 287,further comprising cells cultured on at least one side of thesemi-permeable membrane, wherein the plurality of microchannels arebranched microchannels.